Mitochondria: The Dynamic Organelle (Advances in Biochemistry in Health and Disease, 2) 0387699449, 9780387699448

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Mitochondria

ADVANCES IN BIOCHEMISTRY IN HEALTH AND DISEASE

Series Editor: Naranjan S. Dhalla, PhD, MD (Hon), DSc (Hon) Winnipeg, Manitoba Canada Editorial Board: A. Angel, Toronto, Canada; I. M. C. Dixon, Winnipeg, Canada; L. A. Kirshenbaum, Winnipeg, Canada; Dennis B. McNamara, New Orleans, Louisiana; M. A. Q. Siddiqui, Brooklyn, New York; A. K. Srivastava, Montreal, Canada

Volume 1: S. K. Cheema (ed.), Biochemistry of Atherosclerosis Volume 2: S. W. Schaffer and M-Saadeh Suleiman (eds.), Mitochondria: The Dynamic Organelle

Mitochondria The Dynamic Organelle

Edited by

Stephen W. Schaffer University of South Alabama, Mobile, AL

and

M-Saadeh Suleiman Bristol Royal Infirmary, Bristol, United Kingdom

Stephen W. Schaffer Department of Pharmacology University of South Alabama Mobile, AL 36688-0001 USA [email protected]

ISBN-13: 978-0-387-69944-8

M-Saadeh Suleiman Bristol Royal Infirmary Bristol BS2 8HW UNITED KINGDOM [email protected]

e-ISBN-13: 978-0-387-69945-5

Library of Congress Control Number: 2007921199 © 2007 Springer Science +Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper. 9 8 7 6 5 4 3 2 1 springer.com

Preface

The term mitochondrion is derived from Latin, with mitos meaning thread and chondrion meaning granules. Indeed, under the light microscope, mitochondria often appear as rods or granules within the cytoplasm. For decades after initial visualization of mitochondria by light microscopy, mitochondrial function remained clouded. However, with the development of differential centrifugation and electron microscopy, it was discovered that a chief function of the mitochondria was the generation of ATP for the remainder of the cell. For many years, the energy generating function of the mitochondria was considered the primary, if not the sole function of the mitochondria. During that period, investigators attempted to obtain information on the mechanism of ATP synthesis and the regulation of electron transport. In the first chapter of the book, Dr. Hassinen summarizes those studies, providing clear pictures on the transformation of reducing equivalents into a proton gradient and the mechanism by which the F1 F0 ATPase utilizes the proton gradient to generate ATP. He also summarizes the key regulatory steps of the citric acid cycle, which is the major source of reducing equivalents for the electron transport chain. In the heart, most of the carbon that feeds into the citric acid cycle is derived from fatty acid metabolism. Although fatty acid utilization provides most of the ATP for contraction, a proper balance must be maintained between the utilization of fatty acids and that of glucose. In the second chapter, Drs. Folmes and Lopaschuk discuss the interrelationship between carbohydrate and fatty acid metabolism and how the interrelationship is affected in various disease states. Correction of metabolic dysregulation is introduced as a novel approach toward the treatment of heart disease. In the third chapter, Drs. Sugden and Holmes describe the profound influence of the nuclear regulators, known as the peroxisome proliferator-activated receptors (PPARs), on lipid and carbohydrate metabolism. The physiological and pharmacological ligands that activate the receptors are discussed, along with pathological conditions associated with dysregulation of the receptors. Today, it appears that crosstalk between the mitochondria and the remainder of the cell is an extremely important function of the mitochondria. Not only do the mitochondria provide metabolites to ensure continuation of several metabolic pathways, but they also regulate the levels of key cytosolic constituents. This is made possible by the presence of transporters in the inner mitochondrial membrane that specifically regulate the movement of substrates in and out of the mitochondria. In the fourth chapter, Drs. Kaplan and Mayor describe the structure of a transport protein that shuttles citrate out of the mitochondria, v

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providing a carbon source for fatty acid, triglyceride and cholesterol biosynthesis. The chapter also clarifies the mechanism by which the transporter insures the integrity of the membrane barriers protecting the mitochondria from the accumulation of undesirable substances. In chapter 5, Dr. Vary outlines the role of mitochondria in ammonia detoxification, gluconeogenesis, protein synthesis and acid-base balance. The complex regulation of rate-limiting mitochondrial events, such as flux through pyruvate and branched chain ketoacid dehydrogenases, by transcription factors, hormones and cellular nutrition are reviewed. In chapter 6, Dr. King discusses the important role played by the mitochondria in amino acid metabolism. She reviews evidence that amino acids are involved in mitochondrial-dependent cytoprotective reactions that are dependent on amino acid metabolism rather than protein synthesis. Mitochondria are dynamic organelles. They can migrate, undergo morphological changes and accumulate in response to physiological and pathological biogenetic stimuli. Dr. Al-Mehdi summarizes the dynamic changes induced upon endothelial mitochondria by shear-stress, flow adaptation and flow cessation. Recent studies suggest that the mitochondria may also generate initiators of important signaling pathways. Of particular interest is the activation of cell altering protein kinases by mitochondrial-generated free radicals. How the mitochondria maintain a balance between the generation and destruction of reactive oxygen and nitrogen species is presently unclear. Dr. Turrens argues that it is a complex process, involving multiple sources of reactive oxygen species, several antioxidant enzymes, antioxidant vitamins and an assortment of antioxidant small molecules, all which normally act to prevent the accumulation of damaging levels of oxidants. Like reactive oxygen species, mitochondrial calcium homeostasis must be tightly regulated to ensure mitochondrial and cellular viability. In chapter 9, Drs. Griffiths, Bell, Balaska and Rutter discuss the transporters involved in calcium accumulation by the mitochondria. They also discuss the coupling mechanisms that link modest mitochondrial calcium accumulation with the stimulation of energy production. Also addressed in the chapter are potential pharmacological interventions that might be expected to prevent the damaging effects of excessive mitochondrial calcium accumulation that arise in certain pathological states. Besides the transport of calcium, the mitochondria also regulate the transport of other cations. In chapter 10, Dr. O’Rourke assigns an important cytoprotective role to the potassium channels. He describes experimental evidence showing that multiple potassium channels are cytoprotective, however, he argues that the lack of structure/function relationships has limited the development of ion selective agents with therapeutic significance. Failure to maintain proper levels of calcium and reactive species within the mitochondria alters mitochondrial function, leading to the formation of the mitochondrial permeability transition pore. The formation of the pore renders the mitochondria leaky to organic and inorganic solutes, causing it to lose the ability to generate ATP and to control matrix volume. Drs. Halestrap, Clarke and Khalilin describe the properties of the pore, including the importance of the mitochondrial permeability transition in the depletion of cellular ATP and the loss of cell viability. They point out that events that accompany ischemiareperfusion, such as reactive oxygen species generation, calcium overload,

Preface

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adenine nucleotide depletion, inorganic phosphate accumulation and collapse of the membrane potential, act as initiators or facilitators of pore formation. It follows that effective therapy against ischemia-reperfusion damage requires rapid closing of the mitochondrial permeability transition pore. In chapter 12, Drs. Suleiman and Schaffer review the mechanisms underlying the onset of cellular apoptosis, an orderly type of death that is initiated either by intrinsic, mitochondrial events or by extrinsic stimuli. Because apoptosis is genetically programmed and proceeds along specified pathways, it is highly regulated. Drs. Suleiman and Schaffer describe the effects of several cardioactive agents, including angiotensin II, calcium, reactive species and -adrenergic agonists, on the apoptotic pathway. They maintain that the inhibition of apoptosis might serve as an important therapeutic approach in the treatment of heart disease. Traditionally, apoptosis and necrotic oncosis represent distinct pathways, with only apoptosis considered a highly regulated, genetically programmed pathway. However, in chapter 13, Drs. Murphy and Steenbergen argue that the two forms of cell death share common steps in accidental insults, such as ischemia-reperfusion. They point out that in both forms of cell death, the mitochondria serve as a source of reactive oxygen species and other pro-apoptotic factors, can contribute to ATP depletion and modulate calcium homeostasis. Therefore, they maintain that a better understanding of mitochondrial regulation should provide novel therapeutic approaches for reducing both apoptosis and necrosis. One mechanism of protecting the cell against both forms of death is preconditioning. In chapter 14, Drs. Phillip, Downey and Cohen propose a signaling cascade triggered by Gi coupled receptors that lead to the opening of the KATP channel. They suggest that attenuation of the mitochondrial permeability transition may serve as the end effector of the preconditioning event. In chapter 15, Drs. Shokolenko, LeDoux and Wilson ascribe a key role for mitochondrial DNA damage in apoptosis. They point out that mitochondrial DNA is much more susceptible to oxidative damage than nuclear DNA. Moreover, the synthesis of most mitochondrially expressed peptides is regulated by a single promoter. Since the proteins encoded by the mitochondria are components of electron transport, mitochondrial DNA damage reduces flux through the electron transport and enhances generation of superoxide. Therefore, it is logical to propose that mitochondrial DNA repair, by reversing mitochondrial DNA damage, can rescue cells from excessive superoxide generation and oxidant-induced apoptosis. Recent evidence that mitochondria are directly involved in cell signaling initiated by pro-apoptotic factors, as well as cell survival stimuli, indicates that the status of the mitochondria determines the viability of the cell. Future therapy is likely to take advantage of this central role of the mitochondria to shift the balance in favor of either cell survival or death. Moreover, with the emergence of mitochondria as an important regulator of cell signaling, new interest in the pathophysiology of the mitochondria should yield fascinating results. Stephen W. Schaffer M-Saadeh Suleiman

Contents

Part 1

Mitochondrial Metabolism

1.

Regulation of Mitochondrial Respiration in Heart Muscle . . . . . . . . . . . Ilmo Hassinen

2.

Regulation of Fatty Acid Oxidation of the Heart . . . . . . . . . . . . . . . . . . . . 27 Clifford D. L. Folmes and Gary D. Lopaschuk

3.

Regulation of Mitochondrial Fuel Handling by the Peroxisome Proliferator-Activated Receptors . . . . . . . . . . . . . . . . . 63 Mary C. Sugden and Mark J. Holness

4.

Molecular Structure of the Mitochondrial Citrate Transport Protein. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Ronald S. Kaplan and June A. Mayor

5.

Regulation of Pyruvate and Amino Acid Metabolism . . . . . . . . . . . . . . . 117 Thomas C. Vary, Wiley W. Souba and Cristopher J. Lynch

6.

Amino Acids and the Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Nicola King

Part 2 7.

3

The Dynamic Nature of the Mitochondria

Mechanotransduction of Shear-Stress at the Mitochondria . . . . . . . . . . . 169 Abu-Bakr Al-Mehdi

Part 3

Mitochondria as Initiators of Cell Signaling

8.

Formation of Reactive Oxygen Species in Mitochondria . . . . . . . . . . . . . 185 Julio F. Turrens

9.

Mitochondrial Calcium: Role in the Normal and Ischaemic/ Reperfused Myocardium . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Elinor J. Griffiths, Christopher J. Bell, Dirki Balaska and Guy A. Rutter ix

x

Contents

10.

Mitochondrial Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Brian O’Rourke

Part 4

Mitochondria as Initiators of Cell Death

11.

The Mitochondrial Permeability Transition Pore – from Molecular Mechanism to Reperfusion Injury and Cardioprotection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 Andrew P. Halestrap, Samantha J. Clarke and Igor Khalilin

12.

The Apoptotic Mitochondrial Pathway – Modulators, Interventions and Clinical Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 M-Saadeh Suleiman and Stephen W. Schaffer

13.

The Role of Mitochondria in Necrosis Following Myocardial Ischemia-Reperfusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 Elizabeth Murphy and Charles Steenbergen

Part 5

Mitochondria as Modulators of Cell Death

14.

Mitochondria and Their Role in Ischemia/Reperfusion Injury . . . . . . . . 305 Sebastian Phillip, James M. Downey and Michael V. Cohen

15.

Mitochondrial DNA Damage and Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . 323 Inna N. Shokolenko, Susan P. Ledoux and Glenn L. Wilson

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349

Part 1 Mitochondrial Metabolism

1 Regulation of Mitochondrial Respiration in Heart Muscle Ilmo Hassinen

1.1.

Introduction

Heart muscle with its continual and heavy use of energy is strongly dependent on the most efficient biological energy provider, the process of oxidative phosphorylation, which is responsible for the aerobic conversion and conservation of the combustion energy of fuel substrates to ATP, the universal cellular energy currency. Its key reactions are localized in mitochondria. The myocardial mitochondrion may be regarded as an archetype of its kind and therefore is also a classical experimental model in research on oxidative phosphorylation. Mitochondria are organelles comprised of an outer membrane and inner membrane, enclosing a matrix space. The latter contains the terminal catabolic pathways such as the tricarboxylic acid cycle, fatty acid -oxidation and pyruvate dehydrogenase all of which extract hydrogen and electrons from fuel substrates with the formation of NADH and reduced ubiquinone, a lipid-soluble isoprenoid quinone in the mitochondrial inner membrane. Heart muscle mitochondria are characterized by maximized inner membrane area, achieved by intricate folding, which in electron micrographs appears as a tight packaging of inner membrane invaginations, the cristae. The inner membrane contains the respiratory chain, a series of redox enzymes, which reduce oxygen to water with electrons donated by dehydrogenations in intermediary metabolism. The respiratory enzyme complexes are in principle redox-driven proton pumps, which use the energy derived from oxidationreduction reactions to translocate protons from the matrix space across the inner mitochondrial membrane. This process leads to establishing an electrochemical potential (gradient) of protons across the inner membrane. The electrochemical potential (H +) has two components, an ‘electric’ F·, produced by the membrane potential () and a ‘chemical’ 2.3RT·p H provided by the transmembrane gradient of hydrogen ions. The proton electrochemical potential serves as a primary form of existence of conserved energy, which is utilized by the reversible proton-pumping F1 Fo ATPase to synthesize ATP from ADP and inorganic phosphate. Import of ADP and export of ADP is performed by the ADP/ATP translocase. For the 3

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operation of this (chemiosmotic) principle of energy conservation, the mitochondrial inner membrane must be proton-tight, so that fuel substrates or metabolites can permeate this membrane only by means of various exchange translocators. Like all intracellular organelles surrounded by two membranes, the mitochondria contain their own miniature chromosome, the mitochondrial DNA (mtDNA), which codes for 13 mitochondrial peptides, all having the character of insoluble intrinsic membrane protein. The mutation frequency of mtDNA is one order of magnitude higher than that of nuclear DNA (Zeviani et al. 1998), and these mutations can lead to mitochondrial diseases. MtDNA derangements include both point mutations and deletions. The latter are the cause of a heart disease phenotype, the Kearns-Sayre syndrome characterized by a cardiac conduction defect (Channer et al. 1988) in addition to the symptoms related to skeletal muscle problems.

1.2.

Heart Muscle Mitochondria

In heart muscle most of the copious mitochondria are aligned in interfibrillar rows with a pitch of nearly one per sarcomere. The mitochondria appear as tight packages of inner membrane cristae within the outer membrane. Another population of mitochondria have a subsarcolemmal location and there is some evidence (Riva et al. 2005) of functional differences between these two populations (Manneschi and Federico 1995; Weinstein et al. 1986). The amount of subsarcolemmal mitochondria increases in certain mitochondrial diseases, and the trichrome staining properties of heaps of subsarcolemmal mitochondria gives rise to the histological picture of ‘ragged red fibers’ (Reichmann et al. 1996) characteristic of mitochondriopathies. Since the respiratory chain is composed of intrinsic membrane proteins, maximizing inner membrane area is supportive of a high rate of oxidative phosphorylation. In fact, the protein occupancy of the membrane is so high that there is not much room left for translational, lateral movement of the enzyme complexes in the membrane. Although ‘supercomplexes’ can be isolated with defined methodology (Schagger 2002), the reactions in the ‘respiratory chain’ are probably more dependent on collision frequency between the complexes (Gupte and Hackenbrock 1988) and intramembrane electron carriers than on a structural organization of a chain.

1.3.

The Respiratory Chain

1.3.1. Complex I Complex I catalyses the initiating reaction of the respiratory chain, the reduction of ubiquinone by NADH. The mechanism of complex I action is the least understood of the mitochondrial respiratory complexes. This is due to the extreme

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5

complexity of the enzyme, apparent redundancy of the number of redox centers and their location with respect to the location of its lipid-soluble electron acceptor, ubiquinone in the membrane. In vertebrates, complex I is comprised of 46 dissimilar subunits with a total molecular mass of 900 kDa. Its bacterial counterpart, NDH-1, has only 14 (or 13 due to gene fusion) subunits, all of which are homologues of the subunits of the mammalian enzyme so that the molecular mass is 550 kDa. Seven of these subunits are homologues of the seven mitochondrially coded subunits of the mammalian enzyme, the remaining seven subunits being homologues of the nuclearly coded mammalian complex I subunits. It is becoming evident that these 14 subunits represent the functional core of the mammalian enzyme. The purified enzyme is large enough to be visible in electron micrographs obtained with negative staining. Both mammalian and bacterial enzymes have an L-shaped form (Guenebaut et al. 1998), with an intramembrane arm and a peripheral arm. Resolution of the holoenzyme to subcomplexes (Sazanov et al. 2000) reveals that all of the redox-active centers are located in the peripheral arm of the enzyme, which contains the NADH-binding site, one molecule of flavin mononucleotide (FMN) and eight iron-sulfur centers. The hydrophobic subunits (ND1-ND6 and ND4L) coded in mtDNA reside in the membrane arm of the complex. The ND1 subunit, which reacts with ubiquinone, has no identified redox center. It is also the target of numerous pathogenic mutations, typically resulting in the Leber hereditary ophthalmic neuropathy (LHON). Complex I is a redox-driven proton pump with a stoichiometry of 4H+ /2e− , i.e. four protons per one molecule of NADH oxidized (Honkakoski and Hassinen 1986; Wikstrom 1984). This high H+ /e− ratio cannot be explained with a classical ‘Mitchelian’ redox loop, because the enzyme contains only one hydrogen carrier amongst several electron carriers. Several hypothetical models of complex I function ranging from Q-cycle type mechanisms (Ragan, 1987; Dutton et al., 1998) to reductant-induced oxidation (Brandt 1997) and conformation-driven proton pump (Ohnishi and Salerno 2005) have been proposed. Sequence homologies to Na+ /H+ antiporters (Friedrich and Weiss 1997) or H+ conductors (Hassinen and Vuokila 1993) suggest that the ND2, ND5, ND4 and ND4L subunits may be involved in proton translocation (Kurki et al. 2000; Garofano et al. 2003; Holt et al. 2003; Flemming et al. 2003; Kervinen et al. 2004)

1.3.2. Complex III Complex III (ubiquinol:cytochrome c oxidoreductase) conveys the electrons from reduced ubiquinone to ferricytochrome c, which becomes reduced. The complex is composed of 11–12 dissimilar subunits, and one of these, cytochrome b, is coded in mitochondrial DNA. Cytochrome b contains two heme B groups, namely, a low potential heme (BL −30 mV) and a high potential heme (BH +90 mV). With respect to the mitochondrial inner membrane these hemes are located on opposite sides, one near the inner face and the other near the outer face. The enzyme has also two quinone-binding domains in the vicinity of

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the two heme groups of cytochrome b, and an iron-sulfur protein center reacting with cytochrome c1 , which further reacts with cytochrome c on the outer face of the inner membrane. Complex III is a proton pump with a stoichiometry of 4H+ /2e− . This can be fully explained by the Q-cycle originally proposed by Peter Mitchell (1975), which is based on spectroscopic findings suggestive of the presence of two different cytochromes b. The 11-subunit complex III from bovine heart has been crystallized, and on the basis of x-ray diffraction pattern its 3-D structure is known at 3-Å resolution (Iwata et al., 1998). Its detailed spatial structure fits the Q-cycle mechanism of action.

1.3.3. Complex IV Complex IV (cytochrome c oxidase) catalyses the reduction of oxygen to water by ferricytochrome c with concomitant pumping of protons (Wikström 1984) with a stoichiometry of 2H+ /2e− . Since the electron transport reaction is vectorial and oriented from cytochrome c on the outer face of the membrane to the oxygen reduction site on the inner face of the membrane, a membrane potential (negative inside) is generated. Thus complex IV contributes to the formation of the electrochemical potential by two modes, translocation of electrical charges alone and protons. Complex IV of mammals has 13 dissimilar subunits. Three of them (I-III) are encoded by mtDNA. Bacterial cytochrome c oxidase is composed of only three subunits and these are homologues of the mitochondrially coded ones. Two mitochondrially-coded subunits contain all of the four redox-active groups (two hemes and two copper atoms) of the enzyme (heme A and the binuclear heme A3 -CuB center in subunit I, and CuA in subunit II). The low number of subunits in the bacterial enzyme indicates that these three subunits constitute the functional core of the enzyme, and the other subunits apparently contribute to its stability and regulation. The remaining 10 subunits of the mammalian enzyme are nuclear coded, and at least two of them are involved in the allosteric regulation of the enzyme. Cytochrome c oxidase has been crystallized from a few tissues and species and its 3-D structure has been determined from x-ray diffraction data, e.g. Paracoccus denitrificans (Iwata et al. 1995) and bovine heart (Tsukihara et al. 1996).

1.4.

ATP Synthesis

Complex V (ATP synthetase, F1 Fo -ATPase) is a reversible H+ -ATPase, which pumps protons across a membrane at the expense of ATP hydrolysis. Conversely, it is able to join ADP and inorganic phosphate by extracting water with concomitant formation of ATP. Complex V has three main parts, a membrane-embedded domain Fo , a catalytic domain F1 , and a joining stalk. The F1 domain is a heterohexamer composed

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7

of three -subunits and three -subunits. Pairs of  and  subunits form three catalytic sites, which adopt three different conformations for three partial reactions of ATP synthesis. In the middle of the stalk there is a rod-formed  subunit anchored in the Fo domain that extends to the center of the F1 domain. The b and  subunits also join the F1 and Fo domains and immobilize F1 in relation to Fo . The F1 Fo -ATPase operates in ATP synthesis like a “nanomechanical engine” (Kato-Yamada et al. 1998; Ueno et al. 2005). In the Fo domain there is a ‘rotor’ composed of 9 to 12 c subunits, depending on species. Subunit a of the Fo moiety channels protons into the rotor. Protons are forced through the channel by their electrochemical potential and cause the rotor to rotate. The  subunit rotates with the rotor, and its bent end in the center of the stationary F1 part deforms the - subunits and their catalytic sites in a sequential manner through three functionally different conformations (Oster and Wang 2003). Thus, the electrochemical energy of the mitochondrial inner membrane is transformed through conformational changes to the F1 catalytic sites, which in turn drives ATP synthesis. Three ATP molecules are formed during one turn of the rotor and subunit . The number of protons required for one turn of the rotor is equal to the number of c subunits in it. With 12 c subunits 12 protons are needed for 3 ATP molecules, i.e. the H+ /ATP ratio is 4. The common experimental finding is that three ATP molecules are formed when one molecule of NADH is oxidized by mitochondria. This is in good agreement with the notion that four protons or electrical charges are translocated by each of complexes I, III and IV. The structure of the F1 -ATPase has been determined by x-ray crystallography at 2.8 Å resolution (Abrahams et al. 1994). The proton ATPase/ATP synthase reaction is reversible, which gives rise to problems under conditions in which the electron transport and proton pumping reactions of the respiratory chain are blocked as in ischemia or anoxia. Dissipation of the proton electrochemical potential through proton leakage leaves the H+ -ATPase as the only supporter of (H +). Thus, in ischemia F1 Fo -ATPase enhances the depletion of ATP. However, there exists an endogenous inhibitor peptide IF1 , which binds to F1 Fo -ATPase under conditions of intracellular acidification or decreased mitochondrial membrane potential (Cabezon et al. 2000). There are big species differences in the IF1 content of the myocardium. The concentration is high in the low-beat hearts of larger mammals, whereas in the fast-beat hearts of small rodents the IF1 concentration is low. Ischemia-induced inhibition of F1 Fo ATPase can be demonstrated even in the isolated perfused rat heart (Ylitalo et al. 2001). Ischemic preconditioning, i.e. a brief ischemia/reperfusion, enhances the inhibition of the ATPase during subsequent ischemia (Vuorinen et al. 1995a; Ylitalo et al., 2001; Penna et al. 2004). However, the significance of F1 Fo -ATPase inhibition during the preconditioning process, the most efficient endogenous mechanism of protection against ischemia/reperfusion damage, remains uncertain (Green et al. 1998).

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1.5.

I. Hassinen

Metabolite Translocation in the Inner Membrane.

As far as oxidative phosphorylation and aerobic glycolysis are concerned, the main transport proteins in the mitochondrial inner membrane are ADP/ATP translocase, phosphate translocase, glutamate/aspartate translocase and 2oxoglutarate translocase. The outer membrane is permeable to most smallmolecular weight substrates of the mitochondrion. In creatine kinase (CK)containing tissues such as brain and muscle, a mitochondrial isomorph of the CK exists that is bound to the intermembrane surface of the inner membrane in close structural and functional linkage to the ADP/ATP translocase. In connection of its cytosolic isoenzyme, the CK form a “creatine phosphate cycle” which provides a means of facilitated diffusion of ATP between the mitochondria and the ATP-consuming reactions in myofilaments (Bessman and Geiger 1981; Kupriyanov et al. 1984). The mitochondrial inner membrane is impermeant to NADH and NAD+ , so that the glutamate/aspartate and 2-oxoglutarate translocators (malate-asparate shuttle) are indispensable for transferring reducing equivalents of cytosolic NADH to the mitochondrial matrix under aerobic conditions.

1.6. Mitochondrial Redox Enzymes of Intermediary Metabolism Having Inner Membrane Ubiquinone as Electron Acceptor 1.6.1. Complex II (Succinate:Ubiquinone Oxidoreductase) For historical reasons, succinate dehydrogenase (succinate:ubiquinone oxidoreductase) is commonly listed as a component of the respiratory chain as complex II, although it is not capable of proton pumping. This enzyme of the tricarboxylic acid cycle contains covalently bound flavin-adenine dinucleotide (FAD) as the prosthetic group and three iron-sulfur centers (bi- tri and tetranuclear). It is anchored to the mitochondrial inner membrane by means of two b-type cytochromes, whose role in the electron transfer is unknown, and two ubiquinone-binding proteins. A common textbook error is the classification of succinate dehydrogenase as a flavin-reducing enzyme, although the net reaction catalyzed is: succinate + ubiquinone ↔ fumarate + ubiquinol. None of the subunits of SDH is encoded in mitochondrial DNA (Anderson et al. 1981).

1.6.2. Electron Transfer Flavoprotein:Ubiquinone Oxidoreductase In fatty acid -oxidation the electrons are fed into the respiratory chain via two routes: In the first, fatty acyl-CoA is oxidized to enoyl-CoA by the FADcontaining acyl-CoA oxidase enzyme, which becomes reduced. This is reoxidized

1. Cardiac Mitochondrial Respiration

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by the electron transfer flavoprotein (ETF) (Beckmann and Frerman 1985), which in turn is reoxidized by ETF dehydrogenase (ETF:ubiquinone oxidoreductase), a metalloflavoprotein of the mitochondrial inner membrane (Ruzicka and Beinert 1977). The electrons are channeled to ubiquinone, a substrate of complex III of the respiratory chain. The next -oxidation reaction catalyzed by 3-hydroxyacylCoA dehydrogenase produces NADH, which is oxidized by complex I (for references, see Schulz 1991). Therefore electrons are fed into the respiratory chain via two routes, one of which bypasses complex I, thus lowering the ATP/O ratio.

1.6.3. Glycerol-3-Phosphate Dehydrogenase Two glycerol-3-phosphate dehydrogenases exist. In the cytosol there is an NAD-linked glycerol phosphate dehydrogenase (Ostro, and Fondy 1977), and in the mitochondrial inner membrane there is an FAD-containing flavoenzyme (glycerol-3-phosphate:ubiquinone oxidoreductase), which is linked to the respiratory chain (Klingenberg 1970). The glycerol-3-phosphate-binding site of the enzyme faces the intermembrane compartment and is then accessible to the cytosolic glycerol-3-phosphate, which cannot penetrate the inner mitochondrial membrane. These two enzymes constitute the glycerophosphate shuttle (Houstek et al. 1975), a mechanism of reoxidation of cytosolic NADH during aerobic glycolysis. In heart muscle the mitochondrial content of the glycerol phosphate dehydrogenase flavoenzyme is low, and the glycerophosphate shuttle is not significant in the myocardium, where aerobic reoxidation of cytosolic NADH occurs by the malate-aspartate shuttle (LaNoue and Williamson 1971). The glycerophosphate shuttle is significant in some other tissues such as the -cells of the pancreatic islets, which show a high rate of glucose oxidation (MacDonald 1981).

1.7. Levels of Regulation of Electron Transport in the Mitochondrion The activity of the mitochondrial respiratory chain is geared to the cellular requirements of ATP, but this subtle regulation operates at several levels. The majority of the ATP-consuming reactions occur in the cytosol, such that the translocation of ATP, ADP, Pi and respiratory substrates, although the entry of reducing equivalents into the respiratory chain must also be considered. In fact, a unifying view of the mitochondrial respiratory control in intact cells has not been achieved, and several competing hypotheses have been proposed. The major in vivo regulation has been attributed to the ADP/ATP translocator, the “substrate level” dehydrogenations (Katz et al. 1989; Heineman and Balaban 1990) and the near-equilibria between the ADP/ATP translocator, F1 Fo -ATPase and redox-linked proton pumps (Wilson et al. 1977; Hassinen 1986; Hassinen and Hiltunen 1975). In skeletal muscle, Michaelis-Menten kinetics approximates

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the ADP-dependent respiratory control reasonably well (Chance et al. 1985). Thus, for evaluation of respiratory control an integrating discussion of metabolic control is needed.

1.8. Substrate Level Regulation of Supply of Reducing Equivalents for the Respiratory Chain 1.8.1. Carbohydrate Oxidation Under aerobic conditions the product of glycolysis is pyruvate, which is oxidized in mitochondria by the pyruvate dehydrogenase complex (PDH) to acetyl-CoA, which enters the tricarboxylic acid cycle. The enzyme is interconvertible by means of phosphorylation to an inactive form (PDHi ) and dephosphorylation to an active dephosphorylated form (PDHa ). Free Ca2+ and Mg2+ ions activate the dephosphorylation by a phosphoprotein phosphatase (Midgley et al. 1987). Because mitochondrial ATP is found as a magnesium salt, changes in the mitochondrial ATP concentration influence mitochondrial [Mg2+ ]f (Wieland and Portenhauser 1974; Denton et al. 1975). Thus, the mitochondrial [ATP]/[ADP] ratio becomes a determinant of the [PDHi ]/[PDHa ] ratio. Because the heart rate and contraction force are linked to [Ca2+ ]f transients that follow the action potentials, the time-averaged cytosolic [Ca2+ ]f becomes dependent on the mechanical work output of the heart. Moreover, it has been shown (MacGowan et al. 2001) that the mitochondrial [Ca2+ ]f follows the time-averaged cytosolic [Ca2+ ]f although the transients become filtered out (Leisey et al. 1993). This means that the activation state of PDH is effectively regulated by the mechanical workload of the heart despite the fact that the [ATP]/[ADP] ratio is not markedly influenced by it. The phosphorylation of PDHa to the inactive PDHi is catalyzed by a protein kinase, which is activated by increased acetyl-CoA/CoA and NADH/NAD+ ratios (Ravindran et al. 1996; Peters 2003). Fatty acid oxidation, which is determined by fatty acid availability, increases the mitochondrial acetyl-CoA concentration (Ala-Rämi et al. 2005). Thus, fatty acid oxidation converts PDH to its inactive form and is an efficient suppressor of myocardial carbohydrate oxidation. Also ketone body oxidation elevates the myocardial concentration of acetyl CoA and subdues glucose oxidation. It follows that pyruvate oxidation becomes regulated by the acetyl-CoA/CoA, ATP/ADP, NADH/NAD+ ratios and cardiac mechanical workload.

1.8.2. Fatty Acid Oxidation The regulation of fatty acid oxidation is poorly understood, largely because -oxidation proceeds to completion and intermediate metabolites do not accumulate. Administration of a fatty acid increases cardiac oxygen consumption. This has been attributed to a decoupling effect of fatty acids on mitochondria (Kornberg 1966), a decrease in the P/O ratio related to the by-passing of complex

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one by a portion of the reducing equivalents (see above) or a reduction of NADH and the resulting increase in the thermodynamic driving force of cell respiration (Hassinen et al. 1990). The regulation of fatty acid oxidation is possible at the level of fatty acid uptake into the cells, their import into the mitochondria (via the acylcarnitine/carnitine transporter) and the process of -oxidation itself. For the latter, it is known that the semiquinone form of ETF is a feedback inhibitor of acyl-CoA dehydrogenase (Beckmann and Frerman 1985), NADH inhibits 3-hydroxyacyl-CoA dehydrogenase, and acetyl-CoA inhibits thiolase enzyme (for references, see Schulz 1991). It has been shown in liver mitochondria that an increase in the mitochondrial matrix volume enhances -oxidation. It is also known that matrix volume is affected by calcium. Thus, it is possible that calcium regulates -oxidation, forming a link between ATP-consuming heart muscle contractions and ATPyielding fatty acid oxidation. Moreover, the matrix volume is also affected by mitochondrial pyrophosphate, and mitochondrial pyrophosphatase is inhibited by calcium. However, in heart muscle the pyrophosphate concentration is not affected by metabolic perturbations (Griffiths and Halestrap 1993). When contractility of an isolated perfused heart is adjusted by changes in the extracellular [Ca2+ ] during fatty acid infusion, the alterations in the mitochondrial redox state of flavins (and NADH/NAD+ ) are small compared with changes in oxygen consumption or the free [ATP]·[ADP]/[Pi ] ratio (Vuorinen et al. 1995b). This suggests that the rate of fatty acid oxidation is regulated more by calciuminduced changes in ATP consumption than by direct effects of calcium on -oxidation. In heart muscle, the main determinant of fatty acid oxidation is their availability, and even low concentrations of fatty acids are able to almost totally suppress carbohydrate oxidation as has been recently shown by the 13 C NMR isotopomer analysis methodology (Ala-Rämi et al. 2005). The mitochondrial acylcarnitine/carnitine transferase I is inhibited by malonyl-CoA, an intermediate of fatty acid synthesis. In lipogenic tissues malonyl-CoA is a major regulator of fatty acid oxidation and ketogenesis (McGarry and Foster 1979). Malonyl-CoA is also synthesized by cytosolic acetyl-CoA carboxylase, which is regulated by 5’-AMP-activated protein kinase in heart muscle, where its only metabolic fate is the decarboxylation by the malonyl-CoA enzyme, whose sole role is likely the regulation of fatty acid oxidation (Dyck et al. 1998; Belke et al. 1998).

1.8.3. Tricarboxylic Acid Cycle The mitochondrial tricarboxylic cycle is the most central metabolic pathway with input from carbohydrate, fatty acid and amino acid oxidation. It feeds the respiratory chain by reduction of NAD or ubiquinone. The regulation of tricarboxylic acid (TCA) cycle is somewhat controversial as is the mutual significance of the TCA cycle and respiratory chain in the regulation of cell respiration.

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There are two modes of regulation of the TCA cycle. First, there is flux regulation, but an equally important mode of regulation involves the maintenance of the pool of TCA cycle intermediates. The TCA cycle is designed for oxidation of acetyl-CoA. The net degradation or oxidation of any of the cycle intermediates necessitates their prior export from the cycle. The intermediates act by mass action to accelerate TCA flux but are not consumed by the cycle itself. In spite of that, the total pool size of the TCA cycle in heart muscle undergoes large fluctuations and some of its intermediates function as feedback regulators of cytosolic metabolic pathways. 1.8.3.1.

Flux Regulation

Three enzymes of the TCA cycle have properties suitable for flux regulation. (1) The mitochondrial concentration of oxaloacetate (OxAc) is within the range of the Km of citrate synthase (Siess et al. 1984), such that the reaction velocity becomes dependent on the mitochondrial concentration of OxAc, which in turn depends on the status of the malate dehydrogenase reaction (Forsander 1970). It is known that the activity of malate dehydrogenase is near-equilibrium, such that the concentration of OxAc varies as a function of the mitochondrial NADH/NAD+ ratio, which in turn is dependent on the activity of the respiratory chain. (2) Calcium is an activator (Nichols et al. 1994), NADH a product inhibitor and ADP an allosteric activator (Ehrlich and Colman 1982) of NADdependent isocitrate dehydrogenase, which accounts for the feedback regulation of the enzyme by the respiratory chain. (3) Calcium is an activator of 2-oxoacid dehydrogenase at concentrations encountered in mitochondria (McCormack and Denton 1990). As described above, the mitochondrial [Ca2+ ] follows the time averaged cytosolic calcium. Thus, the activity of 2-oxoacid dehydrogenase is dependent upon the mechanical work output of the heart. 1.8.3.2.

Pool Size Regulation

An important aspect of the TCA cycle is the maintenance of its pool size. Although carbon shuffling between intermediates occurs, TCA cycle flux does not affect total pool size. However, cycle metabolites are also used in some other metabolic routes, biosynthetic for example. Thus, the pool requires continual replenishment, a process referred to ‘anaplerosis’ by Kornberg (1966). It should also be borne in mind that anaplerosis is important not only because it elevates the levels of TCA intermediates but also because it regulates flux through other metabolic pathways. A classical example is citrate, which regulates phosphofructokinase, a rate limiting enzyme in glycolysis. Because the pool size is dependent on both replenishment and disposal, it is also influenced by the metabolic state of the cell. This happens for example during the metabolism of odd-number carbon fatty acids, which produce propionate, which in turn feeds into the TCA cycle via propionyl-CoA carboxylase, thereby producing succinyl-CoA. These TCA cycle disposal mechanisms are sometimes called ‘cataplerotic’ There are several means of anaplerosis in the heart muscle: (1) coupled alanine and aspartate transaminations which produce oxaloacetate, (2) the

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purine nucleotide cycle which by sequential purine deamination and amination produces ammonia and fumarate, (3) glutamate dehydrogenase which produces 2-oxoglutarate, a reaction that is dependent on the mitochondrial NAD+ /NADH ratio, (4) pyruvate carboxylase which produces oxaloacetate, (5) ‘malic’ enzymes (decarboxylating malate dehydrogenases) that produce malate from pyruvate by their reverse reactions. Pyruvate carboxylation in the myocardium is well documented (Peuhkurinen and Hassinen 1982), but its rate during biotin deficiency and the kinetic properties of the malic enzyme suggests that the latter is the main route of pyruvate carboxylation in heart muscle (Sundqvist et al. 1989). Ala-Rämi et al. (2005) have shown that anaplerotic flux is dependent on the work output of the heart. The opposing ‘cataplerotic’ disposal reactions have been less characterized. The redox state of the malic enzyme is such that the net rate is anaplerotic. Only during high rates of anaplerosis can malic enzyme function in the direction of cataperosis (Sundqvist et al. 1987).

1.9.

Respiratory Chain Regulation

1.9.1. Substrate-Level or Respiratory Chain The rate of oxidative phosphorylation and oxygen consumption is adjusted according to the energy needs of the cell, and the phenomenon of “respiratory control” in suspensions as first described by Lardy and Wellman (1952). Nonetheless, one must take into consideration the regulation exerted by the “rate-limiting” steps of mitochondrial electron transport, including (1) control by substrate supply affecting the production of NADH, adjusted by the various regulators of mitochondrial and cytosolic dehydrogenations, (2) control over mitochondrial matrix dehydrogenases, (3) control by the mitochondrial availability of ADP and Pi , (4) control of cytochrome oxidase by the ATP/ADP·Pi ratio through equilibrium reactions in respiratory complexes I and III, (5) allosteric or covalent regulation of cytochrome oxidase, (6) oxygen availability and (7) regulation of expression of the genes coding for the subunits of the respiratory chain complexes.

1.9.2. NADH Limitation The production of NADH or other kinds of reducing equivalents has two important roles in the regulation of cell respiration. First, the oxidation-reduction state of the mitochondrial matrix determines the redox gap and Gibbs free energy change throughout the respiratory chain, ending with the reduction of oxygen. Irreversible thermodynamics predicts that the reaction rate is proportional to the driving force. Second, if the reaction rate of oxygen reduction is determined by the redox state of cytochrome oxidase by means of the reversible redox-driven proton pumps and H+ -ATPase, the set point of the redox state of

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cytochrome oxidase would be indefinite, without the control of the redox state of the NADH/NAD+ couple. A significant amount of evidence implicating an important regulatory role for NADH production has been obtained by means of in vivo phosphorus NMR of the canine heart. It has been shown that during phenylephrine-induced work output transitions the myocardial concentrations of inorganic phosphate, creatine phosphate and ATP remain constant within the accuracy limits of the method (Katz et al. 1989). These data suggest that the adenylate system does not regulate electron transport and oxygen consumption under in vivo conditions, although respiratory control by the adenylate system can be demonstrated in suspensions of isolated mitochondria.

1.9.3. Ca2+ As described above in section 1.8, several possibilities exist for the regulation of mitochondrial NADH production in the myocardium. The main emphasis is on the adjustment of tricarboxylic acid cycle flux by calcium. The NADlinked isocitrate and 2-oxoglutarate dehydrogenases are sensitive to activation by calcium at concentrations observed in the mitochondrial matrix. A correlation between intracellular calcium and oxygen consumption has been observed (Wu et al. 1992), but experiments with the calcium sensitizer levosimendan failed to distinguish between energy consumption or calcium elevation as the cause of enhanced oxygen consumption (Todaka et al. 1996). When the work output of isolated hearts perfused with buffer containing medium chain fatty acids was altered by changes in extracellular calcium, the mitochondrial redox state and cellular energy state data obtained by whole organ fluorometry and 31 P NMR, respectively, supported the notion that oxygen consumption is mainly determined by energy expenditure (Vuorinen et al. 1995b).

1.9.4. Regulation by ATP, ADP and Inorganic Phosphate 1.9.4.1.

Mitochondrial ADP Availability

Oxidative phosphorylation is dependent on mitochondrial availability of ADP and inorganic phosphate, which are transported by the ADP/ATP translocator and phosphate translocator, respectively. The former is electrogenic because of a difference in the net charges of ATP and ADP, and becomes linked to the mitochondrial membrane potential. The phosphate translocator is a phosphate− /OH− exchanger and thus is coupled to transmembrane p H (Palmieri et al. 1993). The ADP/ATP translocase is an abundant protein in mitochondria, but atractyloside administration demonstrates its potential ability to regulate the rate of oxidative phosphorylation at least in isolated mitochondria (Groen et al. 1982). However, a detailed analysis and modeling of skeletal muscle mitochondria (Korzeniewski and Mazat 1996) according to the principles of Kacser (1983) yields a flux control coefficient of 0.14 for the ADP/ATP translocator; the flux

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control coefficient is defined as the ratio of the fractional change of flux to the fractional change in the concentration of the enzyme, with the sum of the coefficients of the pathway being unity (Kacser and Burns 1973). 1.9.4.2.

ADP/ATP Feedback

The signal from the cytosolic energy state can be conveyed to the mitochondria through several modes, such as the concentration of ADP or ATP, the [ATP]/[ADP] ratio, and the [ATP]/[ADP]· Pi ] ratio. In some cases, an index of available high energy phosphate bonds, namely, the “energy charge” (EC) = ½(2·[ATP] + [ADP])/([ATP] + [ADP] + [AMP]), (Atkinson 1968) has been used to express the energy status even though it does not reflect the thermodynamic quantity of free energy available from ATP hydrolysis. The concentration of free ADP in heart muscle cells is very low (30-90 M), compared with the free concentration of ATP (about 5 mM), or inorganic phosphate (around 1 mM), so that during fluctuations within the in vivo range of the ATP/ADP ratio the concentration of ATP remains almost constant. The variation of oxygen consumption upon changes in work output of the isolated working heart preparation (Williamson et al., 1976) has been used to model the metabolic state of the mitochondria/cytosol/myofibril system (Vendelin et al. 2000) relative to oxidative phosphorylation, ADP/ATP translocase, phosphate translocase, mitochondrial and cytosolic creatine kinases and adenylate kinase. It was found that a model assuming equilibrium satisfactorily described the state of ATP, CrP and Cr, but the concentration of ADP changed in a concentration-dependent, non-equilibrium manner indicating the existence of intracellular concentration gradients. The data also revealed a reasonable fit to a Michaelis-Menten type hyperbolic relationship between cytosolic ADP and the rate of oxygen consumption (Vendelin et al. 2000). The extreme sensitivity of the rate of oxidative phosphorylation to ADP, resulting in a nearly constant value for the ATP/ADP ratio is difficult to explain. It has been found that the regulation of oxidative phosphorylation does not follow simple Michaelis-Menten kinetics but is at least second order with respect to ADP (Jeneson et al. 1996). Thus, there must be an amplification mechanism to increase the sensitivity of mitochondria to cytosolic ADP. To test this hypothesis, an analysis was performed using curve fitting which was unbiased toward the specific mechanism of the reactions involved. A satisfactory fit for the dependence of the oxidative phosphorylation rate on ADP concentration was obtained when an arbitrary Hill equation was used, yielding an n value of 3.5. This finding supported the concept that an amplification mechanism sensitizes the mitochondria to ADP (Jeneson et al. 1996) and argues in favor of an allosteric regulatory mechanism. The parallel activation of ATP-consuming and ATP-yielding pathways by calcium has, indeed, been a popular explanation for the relative constancy of the ATP/ADP ratio during workload transitions in heart muscle. Korzeniewsky et al. (2005) have modeled oxidative phosphorylation in different muscles and found that the closest fit to the experimental data is obtained when it is presumed that

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an “each-step activation” occurs. This means that a common signaling molecule influences all contributors of the pathway, i.e. upon activation of muscle ATP consumption, all individual enzyme complexes of the oxidative phosphorylation system become simultaneously activated (Korzeniewski et al. 2005). However, it is difficult to conceive the identity of the messenger. The experimental data indicate that a source of hysteresis exists in the system, such that the activated state of ATP synthesis is not immediately shut off after cessation of ATP consuming muscle stimulation, giving rise to an overshoot of creatine phosphate and undershoot of inorganic phosphate. According to Korzeniewski (1998), the best fit of the experimental data assumes that the enhanced rate of ATP synthesis decays with a time constant of 5 min. The flux control coefficients of the complexes of the oxidative phosphorylation system and the related metabolite translocases have been estimated in skeletal muscle mitochondria, by fitting a theoretical model to the experimental data (Korzeniewski and Mazat 1996). In the resting state (state 4) control was largely imposed by the leakage of protons from the inner membrane, while in the intermediate state (state 3.5), control was largely exerted by the ATP consuming processes themselves. During the maximal rate, rate of oxidative phosphorylation (state 3) the control coefficient of complex III was 0.26 when the ADP/ATP carrier had a coefficient of 0.14 and the ATP synthase 0.16 (Korzeniewski and Mazat 1996). The titration of rat heart mitochondria with oligomycin has yielded a flux control coefficient of 0.04-0.09 for the F1 Fo -ATPase (Ylitalo et al. 2001). 1.9.4.3.

Near-Equilibria in the Respiratory Chain

The classical measurements of the redox potential of the matrix NADH/NAD+ couple and cytochrome c, as well as the free energy change of cytosolic ATP hydrolysis in isolated liver cells have demonstrated that near-equilibrium prevails (Wilson et al., 1974). The same relationship has been found in the isolated perfused rat heart (Hassinen and Hiltunen 1975). An extended network of redox- and free-energy near-equilibria (Hassinen and Hiltunen 1975) prevails in the isolated perfused rat heart (Hassinen 1986), including the redox span across mitochondrial complexes I and III, the ADP/ATP translocase, F1 Fo -ATPase and + H (Kauppinen et al. 1980; Kauppinen et al. 1983; Nuutinen et al. 1981; Nuutinen 1984). Recent data (Liimatta et al. 2004) obtained by organ spectrophotometry of isolated perfused hearts from myoglobin knock-out mice also argue that the dominant regulation exists at the level of the respiratory chain. Mathematical modeling of these equilibria based on the kinetic properties of cytochrome oxidase, which obeys first order kinetics, predicts that oxygen consumption is proportional to the cytosolic [ATP]/[ADP]· Pi ] ratio (Wilson et al. 1977). The equilibria minimize the entropy increase and enhance the energy conservation efficiency of oxidative phosphorylation. This paradigm contradicts simple Michaelis-Menten kinetics for ADP regulation of cell respiration but is in accord with the classical notion that in a metabolic pathway the “rate-limiting” step is out of equilibrium and controls flux through the subsequent steps of the respiratory chain.

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However, it should be borne in mind that a single rate-limiting step does not exist. All steps contribute, although the distribution of control is uneven (Korzeniewski and Mazat 1996). The control attributed to individual steps can be described by their flux control coefficient.

1.9.5. Allosteric and Covalent Regulation of the Respiratory Chain 1.9.5.1.

Complex I

The 42 kD (NDUFA10) subunit of complex I is phosphorylated at some tyrosine residues that remain phosphorylated even during isolation of the complex (Schilling et al., 2005). Moreover, the 18 kD (AQDQ) subunit in the iron-sulfur protein fraction of complex I is phosphorylated in vitro by mitochondrial protein kinase-A (Papa et al. 1996). The 18 kD subunit is coded by the nuclear NDUFS4 gene, and its mutation results in complex I deficiency when homozygous. A 5 bp duplication in the NDUF4 gene destroys the phosphorylation site and abolishes the cAMP-dependent activation of complex I in fibroblast cultures. A deficiency of the 18 kD subunit due to a mutation leading to translation termination a few residues after the mitochondrial targeting sequence results in a defect in the assembly of complex I (Papa et al. 2002). The 18 kD complex I subunit can be dephosphorylated by a mitochondrial serine phosphatase which is inhibited by calcium in contrast to the mitochondrial PDH phosphatase which is activated by calcium (Signorile et al. 2002). Cyclic AMP and PKA have been found in mitochondria, and the A-kinase anchoring proteins (AKAPs) which are variant splicing products of the AKAP1 gene have tag for targeting AKAPs and their PKA complexes to the mitochondrial outer membrane (for a review, see Feliciello et al. 2005). Cytosolic cAMP does not penetrate the mitochondrial inner membrane (Muller and Bandlow 1987), so that its signal probably reaches the mitochondrion by means of a “transductosome” composed of a protein conglomerate assembled by AKAP (Feliciello et al. 2005). 1.9.5.2.

Complex IV

The multiplicity and overlap of the regulatory possibilities of cytochrome oxidase is promoted by the allosteric regulation of mammalian cytochrome oxidase. There are seven high affinity ATP binding sites and ten ADP binding sites in bovine heart cytochrome oxidase. ATP binds to the matrix domain of the subunit IV and acts as an allosteric inhibitor of the enzyme (Arnold and Kadenbach 1997). ATP is also bound to the heart isomorph of the VIa subunit, decreasing the proton pumping efficiency (H+ /e− ratio) of the enzyme (Frank and Kadenbach 1996). On the other hand, ADP bound to the heart type VIa subunit acts as an allosteric activator of the enzyme (Anthony et al. 1993). It is noteworthy that this is specific for the heart enzyme, so that the liver enzyme is not activated by ADP.

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Cytochrome c oxidase is also subject to covalent modification by means of protein kinases. It has been shown that a cyclic AMP dependent protein kinase exists in bovine heart mitochondria that is capable of phosphorylating some mitochondrial proteins (Technikova-Dobrova et al. 2001; Papa et al. 1999). Allosteric inhibition of cytochrome c oxidase by ATP is reversibly switched on by cyclic AMP-dependent protein phosphorylation of subunits II and Vb (Bender and Kadenbach 2000). Also subunit IV is phosphorylated, but the identity of the protein kinase responsible for that is not known (Steenaart and Shore 1997).

Figure 1.1. Metabolic interlinks of mitochondrial respiratory control in cardiomyocytes. Abbreviations: C-I, complex I; C-III, complex III; C-IV, complex IV; C-V, complex V; ANT, ADP/ATP exchange carrier; TCA, tricarboxylic acid; CKm , mitochondrial creatine kinase; CKc , cytosolic creatine kinase; PK-A, cAMP-dependent protein kinase; PLC phospholipase C. The solid arrows show metabolite transport and the dashed arrows depict regulation.

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19

Concluding Remarks

In heart muscle, a tissue of continually high energy consumption, the regulation of electron transfer in the mitochondrial respiratory chain is multifaceted (Figure 1.1). It is evident that several organizational levels contribute to the rate adjustments. The level of dominance is dependent on the stimulus resulting from enhanced ATP consumption. It is apparent that when the primary stimulus is external (humoral) and activates a second messenger system, control is distributed because of the simultaneous activation of substrate-level metabolism, which enhances the production of reducing equivalents to feed into the respiratory chain, and the ATP consuming reactions, which through ADP stimulates the respiratory chain. Under conditions in which the stimulus is non-humoral, control by the adenylate system becomes dominant. The regulation is subtle and the apparent sensitivity of mitochondrial respiration to ADP is extremely high, and some modeling studies indicate that some kind of amplification system exists. This may be based on the simultaneous activation of all of the enzyme complexes of the respiratory chain, although the identity of the messenger remains unknown.

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Wu ST, Kojima S, Parmley WW, Wikman-Coffelt J (1992) Relationship between cytosolic calcium and oxygen consumption in isolated rat hearts. Cell Calcium 13: 235–247 Ylitalo K, Ala-Rämi A, Vuorinen K, Peuhkurinen K, Lepojärvi M, Kaukoranta P, Kiviluoma K, Hassinen IE (2001) Reversible ischemic inhibition of F1 Fo -ATPase in rat and human myocardium. Biochim Biophys Acta 1504: 329–339 Zeviani M, Tiranti V, Piantadosi C (1998) Mitochondrial disorders. Medicine (Baltimore) 77: 59–72

2 Regulation of Fatty Acid Oxidation of the Heart Clifford D. L. Folmes and Gary D. Lopaschuk

2.1.

Introduction

The heart has very high energy demands and requires a large amount of ATP in order to maintain contraction and ionic homeostasis (Opie 1998). To meet this high demand, the heart acts as a metabolic omnivore, metabolizing a variety of carbon substrates, including carbohydrates (glucose, lactate, and pyruvate), fatty acids, and ketone bodies (Neely and Morgan 1974; King and Opie 1998; Stanley et al. 2005). Under normal aerobic conditions, the heart preferentially metabolizes fatty acids, which contribute between 60% and 80% of the required ATP (Lopaschuk et al. 1994a; Stanley and Chandler 2002), with carbohydrates contributing the residual 20% to 40%. While there are many similarities in fatty acid oxidation in various tissues, the regulation of fatty acid oxidation in the heart can differ dramatically from regulation in tissues like liver, skeletal muscle and kidney. This chapter will not attempt to compare these differences, but instead will concentrate on the regulation of fatty acid oxidation specifically in the heart. Although peroxisomal -oxidation also occurs in the heart and may play an important role in providing acetyl-CoA for the production of malonyl-CoA, this chapter will focus on mitochondrial fatty acid oxidation. In addition, fatty acids can also undergo - or -oxidation as well; however the existence of these pathways in the heart has yet to be established. Various disease states are associated with a dysregulation of myocardial metabolism; this chapter will review this dysregulation and will introduce the use of metabolic modulators as a novel pharmacological approach to treating heart disease.

2.1.1. Overview of Myocardial Fatty Acid Metabolism An overview of the pathway for fatty acid oxidation is found in Figure 2.1. The major sources of fatty acids for the myocardium are fatty acids bound to albumin and triglycerides in lipoproteins. Free fatty acids are removed from these sources in the coronary vasculature, and transported across the endothelium into the interstitial space by an as of yet unknown mechanism. 27

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Figure 2.1. Schematic overview of myocardial fatty acid metabolism. ACS, acyl-CoA synethase; CPT-1, carnitine palmitoyltransferase-1: CPT-2, carnitine palmitoyltransferase-2; CT, carnitine acyltranslocase; FABPpm, plasma membrane fatty acid binding protein; FFA, free fatty acid; LPL, lipoprotein lipase.

These fatty acids then enter the cardiomyocytes by either passive diffusion or a carrier-mediated process. These fatty acids are transported in the cytoplasm by fatty acid binding proteins and are activated to long-chain fatty acyl-CoAs, a step required for their further oxidation. The acyl moieties are then transported across the impermeable inner mitochondrial membrane by carnitine dependent transport complex. Once in the mitochondrial matrix acyl-CoAs enter the oxidation spiral and for each successive cycle they are reduced by two carbon units and produce one acetyl-CoA, one NADH and one FADH2 . Acetyl-CoA is further oxidized in the Krebs cycle, which results in the liberation of two CO2 , three NADH, and one FADH2 . These reduced electron carriers enter the electron transport chain, the hydrogens are transferred to oxygen to produce water, and their energy is used for ATP synthesis. The remainder of the chapter will outline the regulation of this process and its dysregulation in disease.

2.2.

Fatty Acid Supply to the Cardiomyocyte

In healthy individuals circulating plasma fatty acid levels can range from 0.2 to 0.8 mM during the course of a day. However, in pathological states which are associated with a metabolic stress, such as diabetes and ischemia,

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circulating fatty acid levels can far exceed these normal levels (>1.0 mM) (Lopaschuk et al. 1994b). Plasma fatty acid concentration is dependent on their release from triacylglycerols (TGs) in the adipocytes, which is dependent on the balance of enzymatic activity of hormone-sensitive lipase for TG degradation and glycerolphosphate acyltransferase for TG synthesis (Lopaschuk et al. 1994a). As the name implies, hormone-sensitive lipase is regulated by hormones, including catecholamines which activate the enzyme, and insulin which inhibits the enzyme. Thus pathological states such as ischemic heart disease which increases -adrenergic stimulation, results in elevated plasma fatty acid concentration. This is in contrary to the fed state, where insulin levels are elevated and fatty acid levels are suppressed. These fluctuations in plasma fatty acid levels are important as the rate of fatty acid uptake into the cardiomyocyte is very dependent on the concentration of nonesterified fatty acids supplied in the coronary vasculature (Bing et al. 1954; Wisneski et al. 1987; Lopaschuk et al. 1994a). However due to the hydrophobic nature of free fatty acids, they are never found in the free form and must be associated with hydrophilic moieties such as proteins, carnitine, or coenzyme A, to increase their solubility in the aqueous environment of the plasma or cytoplasm. In the plasma they are found in their nonesterified form bound to high affinity binding sites on albumin, or contained as TGs within TG-rich lipoproteins, chylomicrons and very-low-density lipoproteins (VLDL) (Richieri and Kleinfeld 1995). Even though the molar concentration of fatty acids in lipoproteins is an order of magnitude greater than that of fatty acids bound to albumin, it is believed that albumin fatty acids are the primary source of energy for the heart (Hamilton and Kamp 1999). However TGs from these lipoproteins cannot cross the plasma membrane, and must first be hydrolyzed by lipoprotein lipase (LPL) to free fatty acids. Studies by Hauton et al. have determined the contribution of fatty acids from albumin, VLDL and chylomicrons to fatty acid oxidation (Hauton et al. 2001). These authors showed that TG from VLDL or chylomicrons could support cardiac function; however they were of secondary importance when in the presence of fatty acid bound to albumin. In addition the presence of free fatty acids suppressed utilization of TG from chylomicrons but not from VLDL. Further studies by this group have shown that TG from chylomicrons are oxidized and contribute to the maintenance of mechanical function, while TG from VLDL is mostly incorporated into tissue lipids (Niu et al. 2004). Cardiac muscle is the tissue with the highest LPL expression, likely due to tissue-specific regulatory elements in the LPL gene promoter region that allows for differential regulation (Enerback and Gimble 1993; Fielding and Frayn 1998). LPL is bound to the luminal side of the capillary endothelial cells and the outer surface of the cardiomyocyte by charge interactions with heparin sulfate, a constituent of the glycocalyx (Merkel et al. 2002a; Augustus et al. 2003; Yagyu et al. 2003; Yokoyama et al. 2004). In vitro observations have shown that both active and inactive LPL can anchor lipoproteins to the cell surface, thereby increasing the efficiency of lipolysis (Merkel et al. 1998a; Merkel et al. 2002b). Mice with a cardiac specific deletion of LPL survive, and exhibit increased circulating TGs, decreased uptake of lipoprotein derived fatty acids and

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a suppressed expression of fatty acid oxidation genes, likely due to a downregulation of a PPAR mediated pathway (Merkel et al. 1998b; Augustus et al. 2004). In addition, transgenic mice with cardiac restricted expression of LPL are able to maintain normal circulating lipid levels, suggesting that the heart is an important site for the homeostasis of serum lipid levels (Levak-Frank et al. 1999; Augustus et al. 2003). Following the hydrolysis of lipoprotein TGs, the free fatty acids bind to albumin, until they are transported to the interstitial space by an unknown concentration gradient dependent mechanism. Uptake of fatty acids into the cardiomyocyte occurs by two mechanisms: passive diffusion and a protein-mediated process (Van der Vusse et al. 2000).

2.3.

Fatty Acid Uptake

2.3.1. Passive Diffusion As fatty acids have low solubility in aqueous environments and have high solubility in lipids, they partition into phospholipid bilayers (the kinetics of which is reviewed elsewhere (Hamilton and Kamp 1999)). Therefore when fatty acids dissociate from albumin, they readily adsorb into the plasma membrane despite being in their ionized form at physiological pH (where their pKa is 4.5) (Hamilton et al. 1994). Movement of ionized long-chain fatty acid (LCFA) anions from one leaflet of the membrane to the other is slow (Gutknecht 1988; Schmider et al. 2000), whereas the movement of non-ionized LCFAs is rapid. This process is thus accelerated due to the fact that the pKa of ionization of LCFAs is 7.6 in the phospholipid bilayer. It has been proposed that this so called “flip-flop” model includes several steps: 1) dissociation of fatty acid from albumin; 2) adsorption of fatty acid to the membrane; 3) protonation; 4) flip-flop within the membrane; and 5) desorption into the cytosolic space. (as reviewed (Hamilton and Kamp 1999; Schaffer 2002)).

2.3.2. Protein Mediated Fatty Acid Uptake Several lines of evidence support the idea of facilitated fatty acid transport. Studies in isolated adipocytes where fatty acids were not esterified, showed that permeation could be distinguished from metabolism, and supply of fatty acids by albumin did not limit uptake (Richieri and Kleinfeld 1995). Fatty acid uptake produced nonlinear saturation kinetics as a function of fatty acid concentration and produced a Km of 7 nM, a value expected for a carrier-mediated process in the range of the physiological concentration of fatty acid (Richieri and Kleinfeld 1995). These results were reproduced in cardiac myocytes by Stremmel and Berk (Stremmel and Berk 1986). Competition for the transporter was observed in competition assays between long chain fatty acids, such as oleate and palmitate (Abumrad et al. 1981; Sorrentino et al. 1996). Another study making use of fluorescent fatty acid derivatives showed that greater than 87% of the fluorescent fatty acids remained unesterified inside the

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cell, therefore the competition could not be explained by metabolism (Storch et al. 1991). Another line of evidence is that fatty acid uptake is sensitive to protein modifying agents, such as phloretin which completely blocks membrane flux of fatty acids, DIDS (4,4’-di-isothiocyanostilbene-2,2’-sulphonate) which produces reversible inhibition of transport and antibodies raised against CD36 inhibited uptake (Abumrad et al. 1981; Abumrad et al. 1984; Harmon et al. 1992). For a more comprehensive review of this evidence see the review by Abumrad et al. (1998).

2.3.3. CD36/Fatty Acid Translocase FAT/CD36 was originally identified as a fatty acid transporter based on its binding of two fatty acid uptake inhibitors, DIDS and sulpho-N-succinimidyl FA esters (Harmon and Abumrad 1993; Abumrad et al. 1998). This 88 kDa transmembrane protein is abundantly expressed in fatty acid metabolizing tissues such as skeletal and cardiac muscle, and appears to be the major fatty acid transporter found in the heart (Vannieuwenhoven et al. 1995; Greenwalt et al. 1995; Pelsers et al. 1999). Evidence for a role of CD36 in fatty acid uptake comes both from experimental models and human studies. Transgenic mice that overexpress FAT/CD36 in muscle have reduced circulating fatty acid levels and fatty acid oxidation is increased only during muscle contraction, suggesting that fatty acid uptake is only accelerated when there is a metabolic demand (Ibrahimi et al. 1999). Conversely, CD36 null mice have increased circulating fatty acids and tissue uptake of an iodinated fatty acid analog, [125 I]BMIPP (15-(p-iodophenyl)-3(R,S)-methylpentadecanoic acid) is reduced in heart, skeletal muscle and adipose tissue (Febbraio et al. 1999; Coburn et al. 2000). If hearts from CD36 knockout mice are perfused as isolated working hearts, a 40-60% reduction in fatty acid oxidation occurs, which is compensated for by an increase in glucose oxidation (Kuang et al. 2004). In addition, humans with mutations in CD36 have lower rates of uptake of [123 I]BMIPP compared to normal people, suggesting that CD36 also partially regulates cardiac fatty acid uptake in humans (Kintaka et al. 2002). Taken together, these results suggest that CD36 is an important regulator of fatty acid uptake in the heart. Electrically stimulated contractions of cardiomyocytes in suspension results in increased fatty acid uptake, and this is completely blocked by sulfo-Nsuccinimidyloleate, a specific inhibitor of CD36 (Luiken et al., 2001a). This increase in flux could be a result of either an increase in sarcolemmal abundance of CD36 or an increase in intrinsic transport activity (Luiken et al. 2004a). In similar studies performed in skeletal muscle, the apparent maximum velocity of transport is increased independent of changes in Km. This finding suggests that the increase in fatty acid uptake is dependent on an increase in total transporter in the sarcolemma (Bonen et al. 2000). Subsequent fractionation of cardiac myocytes showed that under basal conditions CD36 is localized at both the sarcolemma membrane and in intracellular stores (Luiken et al. 2002).

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Contraction or an increase in energy demand was shown to induce a translocation of CD36 from an intracellular pool to the plasma membrane, in a similar mode of regulation as the GLUT-4 transporter (Luiken et al. 2001a; Luiken et al. 2003b; Luiken et al. 2004b). To date several signaling proteins that may play a role in the contraction induced translocation of CD36 have been identified, including AMP-activated protein kinase (AMPK) (Luiken et al. 2003b), protein kinase C   or  or extracellular signal-related kinases (Luiken et al. 2004a). Surprisingly a similar phenomenon of increased CD36 translocation to the sarcolemmal membrane is observed in response to insulin stimulation (Luiken et al., 2001a). Despite these interesting results in isolated cell systems, it has yet to be demonstrated if translocation of CD36 occurs in the intact heart and thus it remains to be determined that translocation of CD36 regulates cardiac fatty acid oxidation under physiological conditions.

2.3.4. Plasma Membrane Fatty Acid Binding Protein The plasma membrane fatty acid binding protein (FABPpm) was originally identified based on its binding of radiolabelled oleate (Stremmel et al. 1985). FABPpm is a 40 kDa protein that is loosely associated with the plasma membrane. The evidence supporting a role of FABPpm in fatty acid transport was initially based on an observation that treatment with an antibody raised against this protein results in a reduction in fatty acid uptake in a variety of cells (Berk et al. 1990; Zhou et al. 1992; Luiken et al. 1997). Additional evidence stems from expression of FABPpm in Xenopus laevis oocytes and 3T3 fibroblasts, which resulted in an increase in fatty acid uptake rates (Zhou et al. 1992; Zhou et al. 1994). A variety of conditions show concomitant changes in FABPpm protein expression and fatty acid utilization, including differentiation of 3T3-L1 cells, fasting, endurance training, diabetes and obesity (Zhou et al. 1992; Turcotte et al. 1997; Kiens et al. 1997; Berk et al. 1997; Luiken et al. 2001b).

2.3.5. Fatty Acid Transport Proteins Fatty acid transport protein (FATP) was discovered as a fatty acid transporter by use of an expression cloning strategy in 2T2-L1 adipocytes (Schaffer and Lodish 1994). Expression of this 63 kDa integral membrane protein in a stable fibroblast cell line results in a 3- to 4-fold increase in long-chain fatty acid transport (Schaffer and Lodish 1994). The cloning of this first FATP led to the discovery of a family of evolutionarily conserved proteins, encoded by at least five different genes in the mouse and displaying different tissue-specific expression patterns (Anderson and Welsh 1990). FATP1 is the major isoform found in heart and skeletal muscle (Hirsch et al. 1998; Binnert et al. 2000; Pohl et al. 2000). However the role of this protein in fatty acid uptake is controversial. Bonen et al. (2002) found an inverse relationship between FATP1 protein expression in the plasma membrane and fatty acid uptake, however this may be due to the use of a poor FATP1 antibody in those studies. At the protein level, FATP1 has 40%

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amino acid homology with very-long chain acyl-CoA synthetase (VLACS), and FATP1 has been reported to be a VLACS (Uchiyama et al. 1996; Kiens et al. 1997; Coe et al. 1999). For a more in depth review of this literature please see a review by Brinkmann et al. (2002). It has yet to be determined whether FATP1 has separate fatty acid uptake and VLACS activities.

2.3.6. Synergistic Action of Transporters It has been proposed that CD36 and FABPpm may cooperate to transport fatty acids across the plasma membrane (Glatz et al. 1997; Glatz and Storch 2001; Bonen et al. 2002). Indirect evidence from studies in giant sarcolemmal vesicles suggest that fatty acid uptake may be inhibited by either sulpho-N-succinimidyl oleate or an anti-FABPpm antiserum (Luiken et al. 1999; Turcotte et al. 2000). However when these agents are applied together they do not result in a further decrease in fatty acid uptake (Luiken et al. 1999).

2.3.7. Very-Low-Density Lipoprotein (VLDL) Receptor The VLDL receptor was originally cloned from a rabbit heart cDNA library and showed striking similarity with the LDL receptor cDNA (Takahashi et al. 1992). The heart is the site with the most abundant VLDL receptor expression and indirect evidence suggests that it may play a role in the uptake of fatty acids. A study in Balb/c mice showed that VLDL receptor expression increases with fasting and they (Kwok et al. 1997) proposed a role of the VLDL receptor in delivery of fatty acids as a source of fuel. Additionally these fasting mice also produced an increase in mRNA of other fatty acid metabolism proteins such as acyl-CoA synthetase and LCAD, and an increase of remnant lipoprotein particles (Kamataki et al. 2002). Studies are still required to elucidate the exact role of the VLDL receptor on cardiac fatty acid metabolism.

2.4.

Cytosolic Transport and Activation of Fatty Acids

2.4.1. Fatty Acid Binding Proteins (FABP) FABPs are the cytoplasmic equivalent of albumin. While albumin transports fatty acids between organs, FABPs are required for the bulk transportation of longchain fatty acids to specific intracellular targets, such as the mitochondrial outer membrane. FABPs are a family of low molecular weight proteins (14–15 kDa) with at least 13 members, with slightly differing amino acid sequence. These protein were first identified as having a high affinity for fatty acids (Ockner et al. 1972), however their role in fatty acid metabolism was not determined until recently. Hearts from mice with a disruption of the gene encoding H-FABPc (the heart isoform of the cytosolic FABP) show a 50% reduction in rates of palmitate uptake. In addition, fatty acid utilization by cardiomyocytes isolated

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from H-FABPc deficient mouse hearts are suppressed by 45% despite the fact that the capacity for fatty acid oxidation in heart homogenates does not differ from wild-type animals (Schaap et al. 1998; Binas et al. 1999). A study of skeletal muscle giant vesicles from H-FABP−/− mice demonstrated a similar reduction in fatty acid uptake, but uptake in vesicles from H-FABP+/− is not impaired, suggesting that H-FABP plays a permissive rather than regulatory role in fatty acid uptake (Luiken et al. 2003a). Evidence suggests that fatty acids are desorbed from the inner leaflet of the sarcolemma by collisional interaction of muscle FABPc with the membrane, which differs from liver FABPc whereby the fatty acids are released into the water layer adjacent to the membrane before they bind to FABPc (Storch and Thumser 2000). It has yet to be determined whether FABPc can physically interact with either fatty acyl-CoA sythetase or with the phospholipids of the outer mitochondrial membrane where this enzyme resides. However this interaction would help to aid in the transport of cytoplasmic fatty acids to the enzyme, which activates the fatty acids before they can undergo fatty acid oxidation in the mitochondria.

2.4.2. Activation of Fatty Acids Fatty acids are activated by esterification to fatty acyl-CoAs by the action of a family of fatty acyl-CoA synthetases (ACS), which differ in their chain length specificity and their subcellular location (Knudsen et al. 1993). The heart isoform of long-chain acyl-CoA synthase is localized to the aqueous cytoplasmic face of the outer mitochondrial membrane (Waku 1992). This conversion to a more reactive CoA thioester is a prerequisite for the subsequent catabolic process. Several lines of evidence suggest that ACS plays a role in the uptake of fatty acids. In adipocytes ACS1 is an integral membrane protein, and it has been shown to localize to the plasma membrane and codistribute with FATP1 (Schaffer 2002). Disruption of yeast orthologs of ACS1, FAA1 and FAA4, results in a restriction of fatty acid uptake and utilization (Faergeman and Knudsen 1997; Choi and Martin 1999). A transgenic mouse model that has increased ACS1 expression in the heart produces marked cardiac TG accumulation, even when serum lipid levels are normal (Chiu et al. 2001). Despite the essential role ACS plays in the catabolism of fatty acids, their regulatory role in fatty acid metabolism has yet to be elucidated. Once activated, fatty acyl-CoAs can bind to another cytoplasmic binding protein, acyl-CoA-binding protein (ACBP). This protein copurifies with FABP and has been identified in a number of tissues including heart (Mikkelsen and Knudsen 1987). It has been suggested that this protein may regulate fatty acid metabolism by the vectorial delivery of fatty acyl moieties to distinct metabolic pathways (Faergeman and Knudsen 1997). In addition, these proteins may buffer the concentration of long-chain acyl-CoA esters, which would relieve the end product inhibition of acetyl-CoA carboxylase (ACC) and adenine nucleotide translocase (Rasmussen et al. 1993). These long-chain fatty acylCoAs can then either be esterified to TGs by glycerolphosphate acyltransferase

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(Coleman et al. 2000) or transported into the mitochondria for subsequent -oxidation (Lopaschuk et al. 1994a).

2.5.

Triacylglycerol as a Source of Fatty Acids

TG stores in the myocardium can be very important endogenous sources of fatty acids for fatty acid oxidation (Crass 1977; Paulson and Crass 1982; Saddik and Lopaschuk 1992). In the unphysiological condition of an isolated heart being perfused in the absence of exogenous fatty acids, the heart can meet 50% of its energy requirements by mobilizing its TG stores. However as concentrations of fatty acids delivered to the heart increase, the contribution of these TG pools to myocardial oxidative metabolism decrease; i.e. during perfusion with a high concentration of fatty acids, TG fatty acids still contribute approximately 11% of the total cardiac ATP production (Crass 1972; Saddik and Lopaschuk 1991). This suggests that myocardial triglycerides are a mobilizable endogenous substrate source due to a constant and rapid turnover of the pool. Approximately 10% of fatty acids taken into the heart are first cycled through the TG pool before they undergo oxidation. This TG turnover can be accelerated by adrenergic stimulation (Crass 1977; Paulson and Crass 1982; Stanley et al. 1997; Goodwin et al. 1998; Goodwin and Taegtmeyer 2000), uncontrolled diabetes (Saddik and Lopaschuk 1994) and during reperfusion of the heart following ischemia (Paulson and Crass 1982; Saddik and Lopaschuk 1992).

2.6. Regulation of Fatty Acid Transport into the Mitochondria 2.6.1. Carnitine Transport System -oxidation of fatty acids occurs primarily in the mitochondria, however a small portion also occurs in the peroxisomes (Schulz 1994; Kunau et al. 1995). Before long-chain acyl-CoAs can undergo oxidation, they first must be transported into the mitochondrial matrix. The inner mitochondrial membrane is not freely permeable to acyl-CoA, most likely to maintain separate cytosolic and mitochondrial CoA pools, therefore a carnitine-dependent transport system is required (Lopaschuk et al. 1994a; Kerner and Hoppel 2000). This system consists of three components: carnitine palmitoyl transferase 1 (CPT1), carnitine acyltranslocase (CT) and carnitine palmitoyl transferase 2 (CPT2), please see Figure 2.2. First CPT1, found on the inner surface of the outer mitochondrial membrane (Murthy and Pande 1987), catalyzes the conversion of long-chain acyl-CoA to long-chain acylcarnitine. Secondly CT exchanges long-chain acylcarnitine for free carnitine across the inner mitochondrial membrane. Lastly, CPT2 regenerates long-chain acyl-CoA in the mitochondrial matrix. Of these three enzymes, CPT1 is the major site of regulation in controlling the uptake of fatty acids into the mitochondria (Kerner and Hoppel 2000).

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Figure 2.2. Regulation of fatty acid transport into the mitochondria. ACC, acetylCoA carboxylase; ACS, acyl-CoA synethase; AMPK, AMP-activated protein kinase; AMPKK, AMP-activated protein kinase kinase; CAT, carnitine acetyltransferase; CPT-1, carnitine palmitoyltransferase-1: CPT-2, carnitine palmitoyltransferase-2; CT, carnitine acyltranslocase; FFA, free fatty acid; MCD, malonyl-CoA decarboxylase.

2.6.2. Malonyl-CoA Regulation of Long-Chain Acyl-CoA Transport into the Mitochondria Fatty acid oxidation is tightly controlled at a number of steps, but the uptake of fatty acyl-CoAs into the mitochondria is arguably one of the most important. CPT1 is the major site of regulation as it is strongly inhibited by malonyl-CoA, a compound that binds on the cytosolic side of the enzyme (McGarry et al. 1983; Lopaschuk et al. 1994a; Zammit et al. 1997; Kerner and Hoppel 2000). The heart expresses two isoforms of CPT1 (CPT1 (82 kDa) and CPT1 (88kDa)) with the 82 kDa isoform predominating. The CPT1 isoform is 30-fold more sensitive to malonyl-CoA inhibition that the CPT1 liver isoform (McGarry et al. 1983; Weis et al. 1994; McGarry and Brown 1997). Therefore, CPT1 activity in the heart is between 10 and 50 times more sensitive to malonyl-CoA than in the liver. This makes malonyl-CoA a key endogenous inhibitor of fatty acid oxidation in the heart, as malonyl-CoA levels are inversely correlated with

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fatty acid oxidation rates. (Saddik et al. 1993; Kudo et al. 1995; Stanley et al. 1996; Hall et al. 1996a; Abu-Elheiga et al. 2001). The half-life of malonyl-CoA in the heart is approximately 1.25 minutes (Reszko et al. 2001), making both the synthesis and degradation of malonyl-CoA important in the control of its steady state level. Malonyl-CoA is synthesized by the carboxylation of acetyl-CoA by acetyl-CoA carboxylase (ACC) (Thampy 1989; Kudo et al. 1995; Hall et al. 1996a; Hall et al. 1996b) and degraded to acetyl-CoA and carbon dioxide in the cytosol and mitochondria by malonyl-CoA decarboxylase (MCD) (Kim and Kolattukudy 1978; Dyck et al. 1998; Saha et al. 2000; Kerner and Hoppel 2002; Dyck et al. 2004)

2.6.3. Acetyl-CoA Carboxylase Regulation of Malonyl-CoA Two different isoform of ACC have been identified, a 265 kDa isoform (ACC265 or ACC) and a 280 kDa isoform (ACC280 or ACC); both isoforms are expressed in the heart, although the prominent form is ACC280 and it appears to contribute most of the heart ACC activity (Bai et al. 1986; Ha et al. 1996; Widmer et al. 1996; Dyck et al. 1999). It has been speculated that ACC280 is directly involved in the regulation of fatty acid oxidation because it is found in tissue with high oxidative capacity and is in close association with the mitochondrial membrane that would allow malonyl-CoA produced to directly inhibit CPT-1 activity (Saddik et al. 1993; Ha et al. 1996; Lee and Kim 1999; Abu-Elheiga et al. 2001). Isolated working heart studies from our laboratory (Saddik et al. 1993; Lopaschuk et al. 1994a; Kudo et al. 1995; Kudo et al. 1996; Makinde et al. 1997) provided direct evidence that ACC is an important regulator of fatty acid oxidation. This has been confirmed in isolated myocytes as well (Awan and Saggerson 1993). ACC activity is dependent on the supply of its substrate, acetyl-CoA, however most of the acetyl-CoA in cardiomyocytes resides in the mitochondria (Idellwenger et al. 1978). This is indirect evidence that acetyl-CoA can be derived from the export of mitochondrial acetyl-CoA as acetylcarnitine by the CT (Lysiak et al. 1986; Saddik et al. 1993) or from citrate via the ATP-citrate lyase reaction (Saha et al. 1997; Poirier et al. 2002). Recently, studies making use of 13 C-labelled substrates and isotopomer analysis of malonyl-CoA provided direct evidence that malonyl-CoA is derived from acetyl-CoA produced by peroxisomal -oxidation (Reszko et al. 2004). This study also suggested that long-chain fatty acids underwent partial oxidation in the peroxisomes, and the released products, C12 and C14 acyl-CoAs, were subsequently oxidized in the mitochondria (Reszko et al. 2004). A more in depth review of the regulation of mammalian ACC has been published (Munday 2002). A major regulator of ACC activity in the hearts and skeletal muscle is AMPK, which phosphorylates three serine residues in ACC265 (serines 79, 1200, and 1215), however only phosphorylation status at ser79 corresponds with inactivation of the enzyme (Davies et al. 1990; Ha et al. 1994). Due to an N-terminal

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extension in ACC280 the equivalent phosphorylation site is ser219 (Scott et al. 2002). AMPK is a heterotrimeric serine/threonine kinase which consists of one catalytic subunit () and two regulatory subunits ( and ); the most prominent heart isoforms are the 2, 2, and 1/2 (Stapleton et al. 1996; Dyck et al. 1996; Gao et al. 1996; Thornton et al. 1998). AMPK was originally discovered as a regulator of fatty acid and cholesterol biosynthesis (Carling et al. 1987; Carling et al. 1989). It is now known that AMPK acts as a key molecular regulator of energy metabolism as it functions as an “energy conservation agent” in the liver by inhibiting anabolic processes when ATP levels are depleted (Carling et al. 1994; Carling et al. 1997). However, in skeletal and heart muscle, its action is to up-regulate fatty acid oxidation during occasions of increased energy demand via the phosphorylation and inactivation of ACC. Thus AMPK is a metabolic sensor in these tissues, which monitors the ratios of AMP/ATP and Cr/PCr, via a mechanism involving the direct allosteric modification of the enzyme by AMP, the activation of an upstream kinase (AMPKK), and the inhibition of protein phosphatase 2C activity (Hawley et al., 1995; Ponticos et al. 1998). Despite the discovery of the identity of two of the AMPKKs, LKB1 (Hawley et al. 2003) and Ca calmodulin dependent kinase kinase (Hurley et al. 2005), it appears that there are additional AMPKKs which mediate the ischemic induced activation of AMPK (Altarejos et al. 2005). Our laboratory has shown that in isolated working rat hearts, not only does ACC copurify with the 2 catalytic subunit of AMPK, it can phosphorylate and inactivate both isoforms of ACC, resulting in an almost complete loss of activity (Kudo et al. 1996; Dyck et al. 1999). Many studies have shown a correlation between increased AMPK activity, decreased ACC activity and increased fatty acid oxidation in working rat hearts (Kudo et al. 1995; Kudo et al. 1996; Makinde et al. 1997; Makinde et al. 1998).

2.6.4. Malonyl-CoA Decarboxylase Regulation of Malonyl-CoA MCD catalyzes the decarboxylation of malonyl-CoA, and thus has been proposed to play an important role in the regulation of myocardial fatty acid oxidation. Generally in situations where MCD activity is elevated, malonyl-CoA content is low, which results in elevated rates of fatty acid oxidation (Dyck et al. 1998; Campbell et al. 2002; Dyck et al. 2004). Despite this correlative data, there is a controversy as to where MCD is localized in the cell. As only cytosolic malonyl-CoA is able to inhibit CPT-1, it would be logical that MCD would reside on or near the mitochondrial membrane, however others have reported it to be localized to the mitochondria, peroxisomes or in the cytosol (Alam and Saggerson 1998; FitzPatrick et al. 1999; Sacksteder et al. 1999; Dyck et al. 2000). This differential localization may be due to two putative translational start sequences (Gao et al. 1999), one of which encodes a protein with a mitochondrial targeting sequence and a peroxisomal targeting sequence (SKL), the second of which codes for a shorter protein that lacks the mitochondrial targeting sequence. Despite this controversy, it still appears that MCD plays an important role in

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the regulation of fatty acid oxidation, and patients with MCD deficiencies have phenotypes consistent with alterations in fatty acid oxidation (Brown et al. 1984; Yano et al. 1997).

2.7.

Regulation of -Oxidation

Once acyl-CoAs have entered the mitochondria they undergo -oxidation, which produces NADH and FADH2 for the electron transport chain and acetyl-CoA for further catabolism in the Krebs cycle. -oxidation requires four different steps as outlined in Figure 2.3, each of which is catalyzed by a specific enzyme with specificity for short-, medium- and long-chain acyl-CoA intermediates (Bing et al. 1954). The acyl-CoA undergoes repeated oxidation via these reactions until only acetyl-CoA units remain. The first step of -oxidation consists of the conversion of acyl-CoA to 2-3-trans-enoyl-CoA in the presence of the FAD, and is catalyzed by acyl-CoA dehydrogenase. Enoyl-CoA hydratase then adds water across the double bond to produce L-3-hydroxyacyl-CoA. In the second dehydrogenation step of the pathway, L-3-hydroxyacyl-CoA dehydrogenase catalyzes the oxidation of L-3-hydroxyacyl-CoA to L-3-ketoacyl-CoA,

Figure 2.3. Schematic of the reactions of -oxidation for even-chain saturated fatty acids.

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with the involvement of NAD+ as a cofactor. The final step of the pathway consists of the splitting of L-3-ketoacyl-CoA at the 2,3-position by 3-ketoacylCoA thiolase, in the presence of CoA, to produce one acetyl-CoA unit and an acyl-CoA that is 2 carbons shorter than the original acyl-CoA. This acyl-CoA feeds back to acyl-CoA dehydrogenase to start another cycle of -oxidation, while acetyl-CoA goes to the Krebs cycle for further oxidation. The oxidation of unsaturated fatty acids require additional enzymes. For odd numbered double bonds, oxidation occurs until a 3-cis-enoyl-CoA intermediate is produced, which is isomerized by 3 2-enoyl-CoA isomerase to 2-transenoyl-CoA, which can be further oxidized. The oxidation of even numbered double bonds requires an additional step. First these fatty acids undergo oxidation until a 2-trans, 4-cis-dienoyl-CoA intermediate is produced. This intermediate is reduced by 2,4-dienoyl-CoA reductase to 3-trans-enoyl-CoA which is isomerized by 3 2-enoyl-CoA isomerase to 2-trans-enoyl-CoA, which can be further oxidized. For a more comprehensive review of -oxidation in the heart please see the reviews by Schulz (Schulz 1991; Schulz 1994). -oxidation is primarily dependent on the workload of the heart, as ratios of NADH/NAD+ and acetyl-CoA/CoA decrease in response to an increase in workload (Oram et al. 1973; Neely et al. 1976). The decrease in acetylCoA/CoA ratio releases the suppression of L-3-ketoacyl-CoA thiolase activity and a decrease in matrix acetyl-CoA also activates acyl-CoA dehydrogenase (Oram et al. 1973). A decrease in workload would result in an increase in L-3ketoacyl-CoA, which feeds back and inhibits acyl-CoA dehydrogenase (Olowe and Schulz 1980). Reports in liver suggest that the NADH/NAD+ ratio regulates L-3-hydroxyacyl-CoA dehydrogenase in the presence of low acetyl-CoA levels, however it is unknown whether this is an important site of regulation in the heart (Latipaa et al. 1986). Alternative methods of feedback regulation may exist in the heart. Elevated levels -oxidation or flux through pyruvate dehydrogenase result in an increase in matrix acetyl-CoA. It has been proposed that this excess acetyl-CoA is transported out of the mitochondria by one of two mechanisms: 1) conversion to acetylcarnitine by carnitine acetyltransferase (CAT), transport across the membrane by CT, and conversion back to acetyl-CoA by a cytosolic CAT (Lysiak et al. 1986; Saddik et al. 1993) or 2) removal of citrate from the mitochondria by the tricarboxylic acid transporter and conversion to acetyl-CoA by the ATPcitrate lyase (Saha et al. 1997; Poirier et al. 2002). Each of these processes increases cytosolic acetyl-CoA levels, which has the potential to accelerate the production of malonyl-CoA, and therefore suppress fatty acid oxidation as shown in Figure 2.2. Recently a mechanism to augment fatty acid oxidation has been proposed that involves mitochondrial thioesterase 1 (MTE1) and uncoupling protein 3 (UCP3) (Himms-Hagen and Harper 2001). MTE1 catalyzes the hydrolysis of fatty acyl-CoA molecules to nonesterified fatty acid anions and free CoA, while UCP3 are classically known for uncoupling ATP synthesis from oxidative metabolism, but may act as a fatty acid anion transporter (Palou et al. 1998; Samec et al. 1998). This hypothesis suggests that in the presence of high rates of

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fatty acid oxidation, free CoA may become limiting for CPT-2 and 3-ketoacylCoA thiolase, and hence could suppress the flux thru -oxidation. Therefore hydrolysis of acyl-CoAs by MTE1 will liberate free CoA to allow for continued -oxidation, and the fatty acid anion generated would be transported out of the mitochondria by UCP3 and be regenerated to acyl-CoA in the cytosol (Stavinoha et al. 2004). However this hypothesis has yet to be examined in the intact heart.

2.8. Interregulation of Carbohydrate and Fatty Acid Metabolism Rates of glucose and fatty acid oxidation are coordinately regulated in the heart, with the rate of fatty acid oxidation being the primary physiological regulator of glucose oxidation. Elevated rates of fatty acid oxidation can inhibit pyruvate dehydrogenase (PDH) activity, the rate limiting enzyme of glucose oxidation. This occurs due to an increase in mitochondrial NADH/NAD+ and acetylCoA/CoA ratios which directly interferes with PDH activity, as well as activating the PDH kinase which phosphorylates and inhibits PDH. In addition, glucose oxidation may regulate fatty acid oxidation via the efflux of acetyl-CoA from the mitochondria and its conversion to malonyl-CoA (as described previously). Therefore inhibition of fatty acid oxidation results in an increase in glucose and lactate uptake, as well as oxidation by lowering mitochondrial acetyl-CoA and NADH levels and relieving the inhibition of PDH, and by decreasing citrate levels and inhibition of phosphofructokinase-1, the rate limiting step of glycolysis. The functional consequences in the switch in substrate usage will be discussed in section 2.11.

2.9. Transcriptional Control of Fatty Acid Oxidation Enzymes The regulation of the expression of proteins that are involved in the myocardial catabolism of fatty acid is not well understood. Except for genes that encode thirteen subunits of the respiratory complexes which are found in circular mitochondria DNA (Scarpulla 2002), all the proteins required for fatty acid uptake and oxidation are encoded by nuclear genes. Recent studies have provided insight that many cardiac genes are regulated by the ligand-activated transcription factors, peroxisome proliferators-activated receptors (PPARs) (Berger and Moller 2002; Huss and Kelly 2004). Three PPAR isoforms have been discovered: PPAR, PPAR/ and PPAR/, and their functional specificity is achieved by isoform-specific tissue distribution, or by differences in ligand affinity or cofactor interactions (Braissant et al. 1996; Escher and Wahli 2000). All of these PPAR isoforms are activated when fatty acids bind, and form heterodimers with retinoid X receptors, and then bind to specific PPAR response elements

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(PPREs) which are located within the promoter regions of many metabolic genes. Binding of fatty acids and eicosanoids increase the activity of the heterodimers, therefore they act as lipid sensors, which increase the facility for fatty acid metabolism in response to greater exposure to fatty acids (Berger and Moller 2002; Huss and Kelly 2004). Additionally PPARs also recruit transcriptional coactivators which are required to initiate target gene transcription, such as PPAR coactivator 1 (PGC-1), which when overexpressed increases the mRNA production of metabolic genes and starts mitochondrial biogenesis (Vega et al. 2000; Lehman et al., 2000; Robyr et al. 2000). Once this complex binds to the PPRE, it increases the rate of transcription of the specific gene that it is bound to by recruitment of histone acetyltransferases, which increases access to the transcriptional start site (Barger and Kelly 2000), as shown in Figure 2.4. PPAR is highly expressed in tissues that have high rates of fatty acid oxidation, such as heart, liver and skeletal muscle. At present the endogenous ligand for PPAR is unknown, however the fibrate class of hyperlipidemic drugs are synthetic PPAR ligands, and they increase the expression of fatty acid oxidation genes and the rates of fatty acid oxidation in cardiomyocytes (Gilde et al. 2003). PPAR regulated genes are involved in most steps of cardiac FA utilization including fatty acid uptake, activation to fatty acyl-CoA, transport into the mitochondria, and mitochondrial ß-oxidation. Evidence from PPAR knockout mouse studies show that fatty acid oxidation enzymes are downregulated and have suppressed fatty acid oxidation (Campbell et al. 2002). Mice with a cardiac specific overexpression of PPAR result in an upregulation of fatty acid utilization enzymes and elevated rates of fatty acid oxidation (Finck et al. 2002; Hopkins et al. 2003). PPAR/ is also expressed in cardiomyocytes and PPAR/ selective ligands can increase mRNA for fatty acid oxidation genes and increase fatty acid oxidation in both neonatal and adult cardiomyocytes (Gilde et al. 2003; Cheng et al. 2004b). In addition cardiomyocyte deletion of PPAR/ downregulates mRNA and protein of fatty acid oxidation genes and reduces catabolism of fatty acids (Cheng et al. 2004a). PPARß activation can also rescue the expression of fatty acid oxidations enzymes that are reduced at baseline in PPAR knockout cardiomyocytes (DeLuca et al. 2000; Cheng et al. 2004b). It appears that PPAR and PPAR/ regulate a similar series of genes, however they are not functionally redundant and PPAR/ cannot completely compensate for PPAR in the knockout mice (Huss and Kelly 2004). PPAR is primarily expressed in adipocytes but is expressed at low levels in other tissues including the vascular wall, skeletal muscle, pancreatic -cell, and heart. PPAR is primarily a regulator of lipid storage and adipose formation, and does not appear to play a role in the regulation of fatty acid oxidation in the heart (Gilde et al. 2003; Lee et al. 2003). However a recent study showed that a cardiac specific knockout of PPAR results in hypertrophic growth; however mRNA or protein expression of fatty acid utilization genes or rates of fatty acid oxidation were not measured in this study (Duan et al. 2005). This would suggest that PPAR plays a role in the heart; however PPAR can also have

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Figure 2.4. Regulation of expression of fatty acid metabolism genes by stimulation of peroxisome proliferators activated receptors. PPAR and PPAR/ directly modify genes in the heart, while most of PPAR effects are extracardiac and occur in either the adipose tissue or skeletal muscle tissue. ACC, acetyl-CoA carboxylase; ACOX1, acyl-CoA oxidase 1; Acrp 30, adipocyte complement-related factor 30; ACS, acylCoA synthetase; FAT, fatty acid transporter; FATP-1, fatty acid transport protein 1; IRS-2, insulin receptor substrate 2; LCAD, long-chain acyl-CoA dehydrogenase; LPL, lipoprotein lipase; MCAD, medium-chain acyl-CoA dehydrogenase; MCD, malonylCoA decarboxylase, mCPT-1, muscle isoform of carnitine palmitoyltransferase-1; MTE1, mitochondrial thioesterase 1; PDK4, pyruvate dehydrogenase kinase 4; PEPCK, phosphoenolpyruvate carboxylase; thiolase, 3-ketoacyl-CoA thiolase; TNF, tumor necrosis factor ; UCP1, uncoupling protein 1; UCP3, uncoupling protein 3; VLCAD, very-long-chain acyl-CoA dehydrogenase.

indirect effects on myocardial fatty acid oxidation via decreasing the fatty acid concentration to which the heart is exposed. For example, the insulin sensitizing agent the thiazolidinediones are PPAR ligands, and their mode of action is to sequester fatty acids in the adipocytes, thus lowering circulating fatty acids and triglycerides (Berger and Moller 2002).

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2.10.

C. D. L. Folmes and G. D. Lopaschuk

Alterations in Fatty Acid Oxidation in Disease

2.10.1. Hypertrophy Induced Changes in Myocardial Fatty Acid Metabolism Cardiac hypertrophy has been associated with reduced pre- and post-ischemic function and is a risk factor for the development of congestive heart failure, myocardial infarction and sudden death (Taegtmeyer and Overturf 1988; Frohlich et al. 1992). Hypertrophied hearts have a reduction in the oxidation of longchain fatty acids, however oxidation of octanoate, a medium chain fatty acid, is not different (el Alaoui-Talibi et al. 1992; Allard et al. 1994). This suggests that mitochondrial -oxidation is not impaired in non-decompensated hypertrophied hearts, but some other component of long-chain fatty acid utilization is impaired. A number of alterations may explain the reduction of longchain fatty acid oxidation in the hypertrophied heart. Spontaneously hypertensive rats have suppressed expression of cytosolic FABP and CD36, which would reduce fatty acid supply to the mitochondria (Aitman et al. 1999; Van der Vusse et al. 2000). In addition, long-chain fatty acid transport into the mitochondria may also be suppressed due to reduced carnitine concentrations in the hypertrophied heart, which could limit carnitine availability for CPT1 (Reibel et al. 1983; Allard et al. 1994; el Alaoui-Talibi et al. 1997). It has also been hypothesized that there is impaired interaction between acyl-CoA synthetase and CPT1 which would also suppress long-chain fatty acid transport into the mitochondria (Christian et al. 1998). Hypertrophied hearts which are showing signs of decompensation (heart failure) also have a reduction of mRNA expression of enzymes required for fatty acid oxidation, including long- and medium-chain acyl-CoA dehydrogenase and long-chain 3-hydroxyacyl-CoA dehydrogenase (Sack et al. 1996; Barger and Kelly 1999). Alterations in fatty acid oxidation in heart failure may share similar characteristics of fatty acid oxidation alterations in hypertrophy. These changes will not be discussed here since the metabolic effect of heart failure has recently been extensively reviewed (Stanley et al. 2005). However, similar to hypertrophy, fatty acid oxidation appears to decrease as the severity of heart failure increases. These alterations in fatty acid utilization in the hypertrophied heart have profound effects on ischemic tolerance, as reviewed in (Sambandam et al. 2002).

2.10.2. Diabetes Induced Changes in Myocardial Fatty Acid Oxidation Dramatic changes in energy metabolism occur in diabetes and are associated with impaired contractile function (Lopaschuk 1996). In uncontrolled diabetes fatty acid oxidation can provide between 90 and 100% of the heart’s energy requirement (Randle 1986; Wall and Lopaschuk 1989). Even if diabetic rat hearts

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are perfused in the absence of fatty acids, glucose oxidation still only provides about 20% of the heart’s total ATP requirement and endogenous triacylglyerol provides a major source of energy. These elevated rates of fatty acid oxidation can be partially explained by a deficiency in insulin signaling resulting in a suppression of glucose uptake. The primary reasons for the elevated rates appear to be an increased level of circulating fatty acids and triglycerides and a modification of the intracellular control of fatty acid oxidation. One site of regulation that appears to be important in diabetes is CPT1. It appears that sensitivity of heart CPT1 to malonyl-CoA does not change (Stanley et al. 1999). However, we have shown in streptozotocin diabetic rats (a model of type 1 diabetes), that AMPK activity is elevated which is associated with a significant decrease in ACC activity (Lopaschuk 1996). In addition MCD activity is significantly increased in the heart (Dyck et al. 1998; Sakamoto et al. 2000). Taken together these results suggest that a reduction in malonyl-CoA inhibition of CPT1 would account for elevated rates of fatty acid oxidation in the diabetic heart. Elevated levels of circulating fatty acids may also result in the activation of PPAR and transcription of fatty acid utilization genes (as review in (Young et al. 2002)). Indeed, transgenic mice with a cardiac specific overexpression of PPAR produce a cardiac phenotype that is similar to the phenotype caused by diabetes (Finck et al., 2002). When these mice were treated with a specific PPAR activator, many fatty acid utilization genes were upregulated including M-CPT1, FATP1, FACS1, CD36, UCP2 and UCP3 (Finck et al. 2002). Additionally PPAR also stimulates the induction of PDK4, which results in the phosphorylation and inhibition of PDH, thus further suppressing glucose oxidation. Overall similar results were also seen in hearts from type 2 diabetic animals as reviewed in (Carley and Severson 2005).

2.10.3. Genetic Deficiencies in Fatty Acid Oxidation The last two decades have seen a steady increase in the identification of a number of mitochondrial defects and associated disease states in patients (Gregersen et al. 2001; Gregersen et al. 2004). Deficiencies in many different fatty acid utilization transporters and enzymes have now been discovered. One of the largest groups of enzymes with heritable mutations are the acyl-CoA dehydrogenases (Gregersen et al. 2004). Generally, defects in the catabolism of medium- or short-chain fatty acids are less likely to produce heart failure or cardiomyopathies than defects in long-chain fatty acid oxidation. One of the most common of these deficiencies is in medium chain acyl-CoA dehydrogenase (MCAD) (Tanaka et al. 1992). Over 90% of these cases involved a homozygous mutation at base pair 985 (A985G) of MCAD, which results in the replacement of a lysine with a glutamate at residue 304 that interferes with tetramer assembly and decreases protein stability (Kelly et al. 1992). Specific defects have also been found in longchain and very long-chain acyl-CoA dehydrogenases (Hale et al. 1985; Strauss et al. 1995). Carnitine deficiencies are usually associated with severe cardiomyopathies and are typically dependent on defects in carnitine transport or carnitine translocase

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(Engel and Angelini 1973). For a more in depth review of fatty acid oxidation defects please see the following reviews (Marin-Garcia and Goldenthal 2002; Gregersen et al. 2004). Although the direct myocardial metabolic effects of these defects have not been determined, it is believed that due to suppressed oxidation of fatty acids, there is an accumulation of intermediary metabolites of fatty acid oxidation, such as long-chain acyl CoAs. These intermediates have detergentlike properties which can modify membrane proteins and lipids, and lead to cardiac arrhythmias and conduction defects (Marin-Garcia and Goldenthal 2002). Additionally fatty acids have also been implicated in the induction of apoptosis (Marin-Garcia and Goldenthal 2002).

2.11. Fatty Acid Oxidation in the Ischemic and Reperfused Heart and Optimization of Fatty Acid Oxidation as a Therapeutic Approach to Treat Ischemic Heart Disease Inadequate supply of oxygen during ischemia results in a reduction in overall oxidative metabolism associated with an impairment of ATP production, the magnitude of which is dependent on the severity of ischemia (Neely and Morgan 1974; Opie 1998). During severe ischemia, the primary source of ATP is glycolytic metabolism of glycogen-derived glucose because coronary blood flow is significantly reduced or even halted; thus substrate and oxygen supply to the heart is suppressed and all oxidative metabolism effectively ceases (Neely and Morgan 1974; Opie 1998). As pyruvate dehydrogenase, the rate limiting enzyme of glucose oxidation, is inhibited, the pyruvate derived from glycolysis is converted to lactate to preserve sufficient NAD+ to sustain flux through glycolysis rather than being metabolized in the mitochondria. This uncoupling of glycolysis from glucose oxidation produces protons from glycolytic ATP hydrolysis, and the inability to remove these protons due to the reduction in coronary blood flow is associated with an intracellular acidosis (Liu et al. 1996a; Liu et al. 1996b; King and Opie 1998; Liu et al. 2002). Homeostasis of normal proton concentrations is dependent on a number of pathways, with two of them involving Na+ -dependent mechanisms: the Na+ -H+ exchanger 1 (NHE1) (Karmazyn et al. 1999) and the Na+ -HCO3 − cotransporter (NBC1, 3, or 4) (Sterling and Casey 2002). During normal aerobic perfusion excess intracellular sodium is removed by the Na+ -K+ ATPase, however during ischemia and reperfusion this transporter is inactivated due to a drop in ATP and intracellular pH. Therefore during ischemia and reperfusion proton extrusion is linked to an intracellular Na+ overload and activation of the reverse mode of the Na+ -Ca2+ exchanger (NCX) (Karmazyn and Moffat 1993), as shown in Figure 2.5. Activation of NCX results in the ischemia-induced Ca2+ overload that is associated with irreversible injury such as apoptosis and necrosis, and reversible injury such as arrhythmias and stunning. This ionic derangement is also detrimental because there is an increased need for ATP to correct these ionic

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imbalances, which shunts ATP away from contractile function and results in a decrease in contractile efficiency. Despite the fact that ATP is quickly replenished due to the rapid recovery of oxygen consumption and TCA cycle activity during reperfusion, mechanical function can only recover once Ca2+ levels have normalized (Lopaschuk et al. 1990; Benzi and Lerch 1992; Liu et al. 1996b; Liu et al. 2002). During reperfusion of previously ischemic hearts, fatty acids can provide over 90% of the myocardium’s ATP requirement (Lopaschuk et al. 1990). This excessive use of fatty acids is due to processes discussed earlier in this review, including an increase in substrate supply due to hormonal stimulation of lipolysis in adipose tissue (Oliver and Opie 1994; Lopaschuk et al. 1994b) and due to a decrease in myocardial malonyl-CoA levels and reduced inhibition of CPT-1 (Kudo et al. 1995; Dyck et al. 1999). The reduction in malonyl-CoA is due to the ischemia-induced activation of AMPK and phosphorylation and inactivation of ACC. MCD activity is preserved during reperfusion; therefore there is a reduction in malonyl-CoA levels, and a stimulation of fatty acid oxidation. These excessive rates of fatty acid oxidation have important consequences on glucose oxidation as these pathways are coordinately regulated (Randle et al. 1963). This phenomenon is of particular importance since elevated rates of fatty acid

Figure 2.5. Metabolic effects of ischemia and reperfusion in the heart. ACS, acyl-CoA synethase; CPT-1, carnitine palmitoyltransferase-1: CPT-2, carnitine palmitoyltransferase-2; CT, carnitine acyltranslocase; FABPpm, plasma membrane fatty acid binding protein, GLUT1/4, glucose transporter 1 or 4; LDH, lactate dehydroganse; NCX, sodium/calcium exchanger; NHE, sodium/hydrogen exchanger; PDH, pyruvate dehydrogenase; PDHK, pyruvate dehydrogenase kinase; PDHP, pyruvate dehydrogenase phosphatase.

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oxidation rise produce an allosteric activation of pyruvate dehydrogenase kinase, which phosphorylates and inactivates the pyruvate dehydrogase complex, thus inhibiting glucose oxidation (Neely and Morgan 1974). Therefore elevated fatty acid oxidation rates during reperfusion of ischemic hearts can exacerbate the uncoupling of glucose metabolism during reperfusion. The use of metabolic modulation to inhibit fatty acid oxidation and stimulate glucose oxidation is a novel therapeutic approach that can be utilized to improve cardiac efficiency (McCormack et al. 1996; Liu et al. 1996a; Kantor et al. 2000; Lopaschuk et al. 2003). Modification of myocardial metabolism is one of the key consequences of ischemia and reperfusion that can contribute to reversible or irreversible ischemia-reperfusion injury. An important therapeutic approach would therefore be to reduce fatty acid oxidation and improve the coupling of glucose metabolism. This would prevent the dysregulation of ion homeostasis and the associated impairment of contractile function. Several different agents have already been tested in experimental models and some are currently being used clinically. These agents can be divided into two classes: those that lower circulating free fatty acids and those that directly modify fatty acid oxidation or glucose oxidation. Agents that lower circulating fatty acids include compounds such as -adrenoceptor antagonists (-blockers), glucose-insulin-potassium (GIK) infusions, and nicotinic acid. The hypothesized modes of action of these compounds is the reduction in circulating free fatty acids, which reduces fatty acid supply to the mitochondria and therefore decrease fatty acid oxidation rates and increase the aerobic disposal of pyruvate. Another approach to lower fatty acid oxidation is to directly inhibit fatty acid oxidation. This approach has been shown with a class of piperazine derivatives, including trimetazidine and ranolazine. Trimetazidine is an inhibitor of long-chain 3-ketoacylcoenzyme A thiolase that reduces fatty acid oxidation and increases glucose oxidation via the Randle cycle (Kantor et al. 2000; Lopaschuk et al. 2003). It is the first widely used antianginal drug with a mechanism of action that can be attributed to the optimization of fatty acid oxidation. Ranolazine is an anti-anginal agent that also inhibits fatty acid oxidation (McCormack et al. 1996). Agents which accelerate glucose oxidation have also been shown to reduce ischemic injury. The prototype of this class, dichloroacetate (DCA), acts by way of the inhibition of PDH kinase, thus ultimately activating PDH and glucose oxidation (Stacpoole 1989). For a more in depth review of metabolic modulation please see these reviews ((Lopaschuk et al. 2002; Folmes et al. 2005).

2.12.

Conclusions

The heart is dependent on the oxidation of fatty acids as a source of ATP for contraction and ion homeostasis. Therefore the use of fatty acids by the heart is a highly regulated process and is dependent on many different processes: substrate

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supply and uptake by the cardiomyocyte, transport and activation in the cytosol, transport into the mitochondria and finally -oxidation of fatty acids. Despite the complexity of this regulation, it ensures a continual supply of acyl-CoA for oxidation. However, the detrimental effects of dysregulation of fatty acid utilization are readily apparent in various forms of disease. As a result the use of pharmacological modulators of myocardial metabolism is an exciting new approach for the treatment of heart disease. Acknowledgements. CDLF is a trainee of the Alberta Heritage Foundation for Medical Research, Canadian Institute of Health Research and Tomorrow’s Research Cardiovascular Health Professionals (TORCH). GDL is a Medical Scientist of the Alberta Heritage Foundation for Medical Research.

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Luiken JJFP, Willems J, Van der Vusse GJ, Glatz JFC (2001a) Electrostimulation enhances FAT/CD36-mediated long-chain fatty acid uptake by isolated rat cardiac myocytes. Am J Physiol Endocrinol Metab 281: E704–E712 Luiken JJFP, Arumugam Y, Dyck DJ, Bell RC, Pelsers MML, Turcotte LP, Tandon NN, Glatz JFC, Bonen A (2001b) Increased rates of fatty acid uptake and plasmalemmal fatty acid transporters in Obese Zucker Rats. J Biol Chem 276: 40567–40573 Luiken JJFP, Koonen DPY, Willems J, Zorzano A, Becker C, Fischer Y, Tandon NN, van der Vusse GJ, Bonen A, Glatz JFC (2002) Insulin stimulates long-chain fatty acid utilization by rat cardiac myocytes through cellular redistribution of FAT/CD36. Diabetes 51: 3113–3119 Luiken JJFP, Turcotte LP, Bonen A (1999) Protein-mediated palmitate uptake and expression of fatty acid transport proteins in heart giant vesicles. J Lipid Res 40, 1007–1016 Lysiak W, Toth PP, Suelter CH, Bieber LL (1986) Quantitation of the efflux of acylcarnitines from rat-heart, brain, and liver-mitochondria. J Biol Chem 261: 3698–3703 Makinde AO, Gamble J, Lopaschuk GD (1997) Upregulation of 5’-AMP-activated protein kinase is responsible for the increase in myocardial fatty acid oxidation rates following birth in the newborn rabbit. Circ Res 80: 482–489 Makinde AO, Kantor PF, Lopaschuk GD (1998) Maturation of fatty acid and carbohydrate metabolism in the newborn heart. Mol Cell Biochem 188: 49–56 Marin-Garcia J, Goldenthal MJ (2002) Fatty acid metabolism in cardiac failure: biochemical, genetic and cellular analysis. Cardiovasc Res 54: 516–527 McCormack JG, Barr RL, Wolff AA, Lopaschuk GD (1996) Ranolazine stimulates glucose oxidation in normoxic, ischemic, and reperfused ischemic rat hearts. Circulation 93: 135–42 McGarry JD, Brown NF (1997) The mitochondrial carnitine palmitoyltransferase system. From concept to molecular analysis. FEBS Journal 244: 1–14 McGarry JD, Mills SE, Long CS, Foster DW (1983) Observations on the affinity for carnitine, and malonyl-CoA sensitivity, of carnitine palmitoyltransferase-I in animal and human-tissues - Demonstration of the presence of malonyl-CoA in non-hepatic tissues of the rat. Biochem J 214: 21–28 Merkel M, Eckel RH, Goldberg IJ (2002a) Lipoprotein lipase: genetics, lipid uptake, and regulation. J Lipid Res 43: 1997–2006 Merkel M, Heeren J, Dudeck W, Rinninger F, Radner H, Breslow JL, Goldberg IJ, Zechner R, Greten H (2002b) Inactive lipoprotein lipase (LPL) alone increases selective cholesterol ester uptake in vivo, whereas in the presence of active LPL it also increases triglyceride hydrolysis and whole particle lipoprotein uptake. J Biol Chem 277: 7405–7411 Merkel M, Kako Y, Radner H, Cho IS, Ramasamy R, Brunzell JD, Goldberg IJ, Breslow JL (1998a) Catalytically inactive lipoprotein lipase expression in muscle of transgenic mice increases very low density lipoprotein uptake: Direct evidence that lipoprotein lipase bridging occurs in vivo. Proc Natl Acad Sci USA 95: 13841–13846 Merkel M, Weinstock PH, Chajek-Shaul T, Radner H, Yin BY, Breslow JL, Goldberg IJ (1998b) Lipoprotein lipase expression exclusively in liver - A mouse model for metabolism in the neonatal period and during cachexia. J Clin Invest 102: 893–901 Mikkelsen J, Knudsen J (1987) Acyl-CoA-binding protein from cow. Binding characteristics and cellular and tissue distribution. Biochem J 248: 709–714 Munday MR (2002) Regulation of mammalian acetyl-CoA carboxylase. Biochem Soc Trans 30: 1059–1064

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3 Regulation of Mitochondrial Fuel Handling by the Peroxisome Proliferator-Activated Receptors Mary C. Sugden and Mark J. Holness

3.1.

Introduction

Within the body, lipids are essential for many aspects of cellular processes, and the PPARS are sensors of lipids that act to regulate gene expression. As lipid metabolism and glucose homeostasis are intrinsically related, these transcription factors exert a profound influence on the selection of metabolic fuels for ATP generation via mitochondrial oxidation. This chapter therefore focuses on the role of the PPARs as key coordinators of the selection of metabolic fuels used for mitochondrial ATP production. In man, the nuclear hormone receptor family at present consists of 48 mammalian transcription factors that are related in terms of evolution, function and structure. These transcription factors regulate a variety of genes that control development, inflammation and nutrient metabolism in a range of tissues and organs (reviewed in Berkenstam and Gustafsson 2005), and can be divided into 3 groups. The first consists of classical high-affinity receptors for steroid and thyroid hormones. The second group consists of low-affinity receptors that bind non-classical ligands, including a wide range of metabolic intermediates including fatty acids (FA) and cholesterol derivatives, and are implicated in the regulation of lipid metabolism. The third group consists of “orphan” receptors, such as the estrogen-related receptor  (ERR), for which neither endogenous nor synthetic ligands have yet been identified. The low-affinity nuclear receptors include the peroxisome proliferatoractivated receptors (PPARs). Other important low-affinity receptors include the RAR-related orphan receptors (which bind cholesterol or retinoic acid), the Liver X receptor (LXR) subfamily (LXR and LXR), which are activated by oxidised cholesterol derivatives (oxysterols), and the Farnesoid X receptor (FXR), which binds bile acids. The retinoid X receptor (RXR) is a common binding partner for 63

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the PPARs. LXRs regulate cholesterol homeostasis in liver and adipose tissue and “reverse cholesterol transport” from peripheral tissues to liver, and are also increasingly recognized to be involved in inflammation and atherosclerosis. FXR regulates hepatic intracellular bile acid and cholesterol levels by regulating the expression of genes involved in cholesterol catabolism and biliary cholesterol secretion. Within the body, lipids are essential for many aspects of cellular processes, including oxidative substrates for ATP production and, as such, are integral to mitochondrial function. The PPARs act as lipid sensors. Through the actions of lipids - or their derivatives - to activate the PPARs, lipids influence their metabolic fates in an interactive manner which is fundamental to metabolic fuel homeostasis under conditions of variable lipid availability - either dietary or released from stored triglyceride (TAG). As lipid metabolism and glucose homeostasis are intrinsically related, as initially described by the pioneering work of Randle and his colleagues (Randle et al. 1963); reviewed in Randle (1998), these transcription factors exert a profound influence on not only lipid, but also glucose handling. The PPARs and their roles as key coordinators of metabolic fuel oxidation will form the major focus of this review. The roles of the PPARs in atherosclerosis and inflammation are described in detail elsewhere (Israelian-Konaraki and Reaven 2005; Staels 2005).

3.2. PPARs: General Function in Relation to Tissue Distribution PPAR (also known as NR1C1) was originally identified as a target for hepatocarcinogens that elicited peroxisome proliferation in the liver of rodents (Issemann and Green, 1990). These effects are not observed in man. Subsequent studies identified two further PPAR receptor isoforms: PPAR (also known as PPAR, NR1C2 or FA-activated receptor, FAAR) and PPAR (also known as NR1CR3). Subsequently, PPARs were demonstrated to be closely implicated in mediating adaptive metabolic responses to changes in systemic metabolic fuel availability, in particular lipids (reviewed in Kliewer et al. 2001). More recently, the involvement of PPARs has been identified in a diverse range of physiological and pathological processes. The differing biological actions of the PPAR isoforms reflect differential tissue expression patterns. PPAR, a lipo-oxidative transcription factor, is most abundantly expressed in a range of tissues with a high capacity for FA oxidation and which have a high energy demand related to their physiological functions: these include cardiac myocytes, hepatocytes, kidney cortex and pancreatic islets (Desvergne and Wahli 1999; Kersten et al. 1999; Sugden et al. 2001). PPAR is also present in monocytes/macrophages, the artery wall, vascular smooth muscle cells and endothelial cells (Duval et al. 2002). PPAR promotes a programme of lipid-induced activation of genes encoding proteins involved in FA uptake, -oxidation, FA transport into peroxisomes and -oxidation of unsaturated FAs

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(Braissant et al. 1996; Wolfrum et al. 2001) and is established to be critical to adaptations to fasting. PPAR, a lipogenic transcription factor, has a more limited tissue distribution. It is highly expressed in white and brown adipocytes, the placenta, cells of the immune system (macrophages, monocytes), and the endocrine pancreas (Braissant et al. 1996; Dubois et al. 2000; Lupi et al. 2004; Zhou et al. 1998). PPAR promotes adipocyte differentiation, regulates intracellular lipid accumulation, induces genes involved in lipogenesis (Rosen et al. 1999), alters the secretion of adipokines, and regulates genes involved in or enhancing insulin signalling (Gray et al. 2005; Mueller et al. 2002). Thus, while PPAR does not have a direct effect on mitochondrial fuel selection, it influences partitioning of available dietary lipid between adipose tissue and oxidative tissues. Altered lipid delivery to oxidative tissues such as heart, liver and oxidative skeletal muscle has two effects: it influences competition between FA and glucose as oxidative fuel by virtue of their relative availability and also may influence insulin action within non-adipose tissues should lipid delivery be excessive. In muscle, the strong relationship between PPAR and altered lipid handling is accompanied by increased intracellular concentrations of lipid metabolites, such as acyl-CoA and diacylglycerol, which appear to impair insulin signalling by activating a serine kinase cascade. This culminates in serine phosphorylation of components of the insulin signalling pathway and a resultant impairment of their activation by tyrosine phosphorylation (see e.g. Roden et al. 1996). PPAR is expressed ubiquitously, but is found only at low levels in liver. Of the three PPAR isoforms, the function of PPAR is the least well defined, except perhaps in skeletal muscle. Emerging research has demonstrated distinct tissue-specific patterns of regulation of FA utilisation by PPAR, with a common theme being the promotion of oxidative metabolism (Fredenrich and Grimaldi 2005; Grimaldi 2005; Lee et al. 2003; Luquet et al. 2005). Thus, PPAR acts as a central regulator of FA breakdown, as well as influencing the intrinsic oxidative capability of skeletal muscle.

3.3. PPARs: Domain Structure and Regulation of Transcriptional Activity As with all nuclear receptors, the PPARs have a modular structure comprising a ligand-independent transcriptional activation domain (AF-1) in the N-terminal region, zinc finger motifs, a hinge region important for cofactor docking, a conserved C-terminal ligand binding domain and an AF2 domain that binds to co-repressors and co-activators (Figure 3.1). Each of the PPAR subtypes bind to DNA as obligate heterodimers with a second nuclear receptor, RXR (Kliewer et al. 1992) (Figure 3.1). In their inactive state (in the absence of ligand), the PPARs can repress transcription by recruiting accessory proteins (co-repressors) and recruiting histone deacetylase. Activation

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co-repressor

A

PPAR

RXR

chromatin

B

co-activator ligand

e.g. PGC-1α AF2 AF1

PPRE

target gene

Figure 3.1. Interaction between PPARs, RXRs, co-repressors and co-activators in relation to the regulation of gene expression Binding of the co-repressor to the PPAR/RXR dimer in the absence of ligand blocks gene expression (panel A). Ligand binding leads to a conformational change promoting release of the co-repressor and binding of the co-activator resulting in gene transcription (panel B).

through ligand binding modifies local chromatin structure, inducing a conformational change leading to dissociation of co-repressors and recruitment of protein complexes containing histone acetyl transferase activity that enhance gene transcription (co-activators) (Figure 3.1). Several transcriptional coactivators, including PPAR coactivator-1 (PGC-1), the biological importance of which is discussed later in this review, bind and enhance PPAR activity (Puigserver and Spiegelman 2003). The RXRs can be activated by 9-cis retinoic acid, an endogenous vitamin A derivative. RXR heterodimers formed with PPARs can be activated by RXR and PPAR agonists either independently or together, in which case they elicit

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synergistic activation. The individual roles of each of the RXR isoforms (  ) (Bastien and Rochette-Egly 2004) which complex with the PPARs to form the heterodimeric PPAR:RXR complex has not yet been determined (Berger and Moller 2002). However, like the PPARs, the RXR isoforms are characterised by a tissue-specific distribution: in rat heart, RXR and RXR are most abundant, whereas RXR is only marginally detected (Mangelsdorf et al. 1992). The preferred DNA-binding site, referred to as the DR-1 motif, is the same for each of the PPAR/RXR heterodimer pairs (Kliewer et al. 2001). Specificity of function may be conferred by mechanisms that impart subtype-specific regulation of PPAR activity, including differences in binding affinity for endogenous ligands (Forman et al. 1997; Lin et al. 1999) or through phosphorylation of their transcriptional activation domains (Barger et al. 2000; Barger et al. 2001; Forman et al. 1997; Lazennec et al. 2000). PPAR contains a consensus mitogen-activated protein kinase phosphorylation site which, when phosphorylated, decreases the ligand binding affinity of PPAR (Shao et al. 1998) and negatively regulates transcriptional activity (Adams et al., 1997; Camp et al. 1999; Hu et al. 1996; Zhang et al. 1996). Docking of coactivators is generally regulated by a conformational change. Some of the most interesting of the transcriptional coactivators include PGC-1 and PGC-2. PGC-1, in particular, is a key regulator of cellular energy metabolism: it is enriched in metabolically active tissues including heart (reviewed in Finck et al. 2005). The activated PPAR:RXR complexes bind to specific response elements (PPREs) in the promoters of the various target genes. These are comprised of a consensus sequence (AGGTCA), the NR hexameric core recognition motif, that can be configured as a single element or two tandem repeats separated by one nucleotide. Binding to the PPREs modifies transcription, leading to an altered complement of enzymes in the cell, with a resultant alteration in cellular functional characteristics. Through this mechanism, each PPAR can, once activated, confer distinct properties of lipid handling to the tissues in which it is found, with an overall systemic lipid-lowering effect.

3.4.

Pharmacological PPAR Ligands

The importance of the nuclear hormone receptors in general was revealed when it was discovered that they represented the targets of a number of widely-prescribed drugs. It was discovered that PPAR and PPAR are the molecular targets for major classes of drugs used to ameliorate atherogenic hypertriglyceridemia and insulin resistance, both of which are commonly used in the treatment of type 2 diabetes. The fibrate class of drugs (clofibrate, gemfibrozil, fenofibrate, bezafibrate and ciprofibrate) have been in clinical use for over 40 years. These drugs target PPAR. Synthetic ligands for PPAR, in addition to the fibrate drugs, include phthalate ester plasticers, herbicides, and leukotriene D4 receptor antagonists. PPAR is the molecular target for the insulin-sensitising thiazolidinediones (TZDs) (e.g., rosiglitazone, pioglitazone). Some of the side effects

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of the TZDs include weight gain through effects of PPAR on adipose tissue (described below) and current research is evaluating newer PPAR ligands, whose mode of binding differs from the TZDs or which are selected on the basis of attenuated regulation of lipogenic genes (Shi et al. 2005). A new class of novel PPAR-targeted compounds termed dual-acting PPAR agonists, which include the compound ragaglitizar, specifically activate both PPAR and PPAR. Synthetic PPAR ligands include GW501516 and GW0742, which activate PPAR with a 1000-fold selectivity over other PPAR isotypes (Sznaidman et al. 2003). There is also current pharmaceutical interest in PPAR-pan agonists (which target all three PPAR isoforms): these generate enhanced insulin sensitivity, mild suppression of circulating triglycerides, reductions in low-density lipoprotein (LDL) cholesterol, and increased high density lipoprotein (HDL) cholesterol. A recent review has described emerging concepts in the use of PPAR agonists for treating metabolic and cardiovascular disease (Staels and Fruchart 2005).

3.5.

Physiological PPAR Ligands

The PPARs are vitally important sensors of the lipid status of the body and of individual tissues or organs of the body. Although the complete range of physiological ligands that regulate the activities of each of the PPAR isoforms still remains to be established, it is now known that the PPARs are activated by binding of naturally-occurring endogenous or dietary FA or lipid derivatives. Known natural ligands of PPAR include polyunsaturated FAs and FA derivatives, including eicosanoids and leukotriene B4 (a lipoxygenase product of arachidonic acid). Recently the -hydroxylated derivatives of 14,15-epoxyeicosatrienoic acid, another derivative of arachidonic acid, has been demonstrated to activate PPAR (Cowart et al. 2002): other EETs and their derivatives may also function as other natural ligands. Oleoylthanolamide, a naturally occurring lipid that is structurally similar to the endogenous cannabinoid anandamide, also activates PPAR (Fu et al., 2003). New data also indicate that, in liver, newly-synthesised fat (but not pre-existing fat) may be an endogenous PPAR activator (Chakravarthy et al. 2005). These studies used a mouse model with liver-specific inactivation of the gene encoding the lipogenic enzyme FA synthase (FAS). When normal chow was substituted by a high carbohydrate-zero fat diet, these mice showed a fasting metabolic profile (hypoglycemia, high FA, lowered insulin, low leptin), and a fatty liver: this resembles the phenotype of fasting PPAR null mice. This phenotype was reversed by the provision of highfat diet. Activation of PPAR also “rescued” the phenotype. This suggests that in the absence of dietary fat, FA synthesised de novo through FAS (“new fat”) can act as a PPAR ligand (and upregulate FA oxidation). It was also suggested that dietary FA transported to the liver via chylomicrons could represent another source of “new” fat. However, it was proposed that FA released from stored TAG in adipose tissue, “old fat”, could not activate hepatic PPAR as effectively. It was proposed that new fat and old fat access separate intracellular compartments, each containing a different pool of PPAR, and these pools cannot be

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accessed by the FA derived from the bulk of intrahepatic TAG (most of which originate from adipose-tissue derived FA). One pool of PPAR is activated by both newly synthesised and dietary FA (and upregulates glucose synthesis and FA oxidation); the other pool is activated only by biosynthetic FA, and this pool additionally activates cholesterol balance. The possibility was suggested that a lipid-binding protein specific for new fat shuttles FA to nuclear PPAR or that PPAR shuttles between the nucleus and cytoplasm via chaperones (e.g. hsp90) that promote an interaction with new fat. VLDLs produced by the liver and chylomicrons containing dietary TAG are targeted to heart and skeletal muscle, generating a local supply of FA and monoacylglycerol via lipoprotein lipase (LPL). Physiological TAG hydrolysis mediated by LPL might represent a link between lipid metabolism and PPAR activation in tissues, such as heart, where there is less de novo fat synthesis than in liver a finding relevant to the effects of provision of high-fat diets, particularly those rich in very-long-chain FAs and polyunsaturated FAs, which can induce FA oxidation via PPAR. Lipolysis of TAG-rich lipoproteins (VLDL) generates PPAR ligands in endothelial cells (Ziouzenkova et al. 2003). Cardiac specific knockout of LPL elevates plasma and cardiac TAG and lowers expression of the PPAR target gene pyruvate dehydrogenase kinase (PDK) 4 (Augustus et al. 2004). Whether or not LPL activity in adipose tissue generates natural agonists for PPAR has not been investigated, but it is known that PPAR is activated by specific prostanoids, such as 15-deoxy-1214 prostaglandin J2. Activation of PPAR by components of oxidised LDL (9-hydroxyoctadecadienoic acid, 13- hydroxyoctadecadienoic acid) enhances macrophage lipid accumulation through induction of a scavenger receptor/transporter (FA translocase/CD36) (Lee et al. 2004). PPAR is activated by long-chain FA, both saturated and unsaturated, and by prostacyclin produced by the conversion of polyunsaturated FAs (Amri et al. 1995). Again, it is not yet known whether PPAR activation is coupled with LPL activity in oxidative skeletal muscle.

3.6. The Role of PPAR in the Regulation of Fuel Handling The actions of PPAR in heart and liver reflect the biological roles of these tissues: the heart as a continually-contracting pump, the liver as a major fuel producer and transformer. A recent study has demonstrated that, during neonatal development of the heart, all three PPAR isoforms are upregulated almost in parallel (Steinmetz et al. 2005). In parallel, cardiac metabolism switches from glucose to lipid as the preferred energy substrate to generate ATP (Makinde et al.1998). This transition is accompanied by changes in the activity and expression levels of several enzymes and regulators involved in FA metabolism, indicating a functional bias towards the lipo-oxidative PPARs, namely PPARs - and -.

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3.6.1. Cardiac Fuel Utilisation In the adult, the heart uses most available fuels to sustain a continuous and enormous demand for energy to fulfill its function (Stanley and Chandler 2002). Although pyruvate derived from either glycolysis or lactate that enters the cell via the monocarboxylate transporter (MCT) can be converted to acetyl-CoA via the pyruvate dehydrogenase (PDH) complex for entry into the tricarboxylic acid (TCA) cycle (Figure 3.2), lipids are thought to constitute the predominant energy substrate for the adult heart, even in the well-fed state. FAs account for 70% of total ATP production in the normal heart; this proportion increases to >90% in

VLDL-TAG lipoprotein lipase lactate MCT

FA

glucose GLU

lactate

plasma membrane

CD36

glucose

FA acyl-CoA synthetase

glycolysis pyruvate

endogenous TAG

FA-CoA

mCPT1

PDC

CPT2 PDK4 acetyl-CoA TCA cycle

β-oxidation

FA-CoA

NADH/FADH2

electron transport chain

ATP

Figure 3.2. Regulation of cardiac glucose and lipid utilisation by PPAR. Sites regulated by PPAR are shown in bold. Abbreviations: AQP, aquaporin; CPT, carnitine palmitoyltransferase; GLUT, glucose transporter; FA, fatty acid; LPL, lipoprotein lipase; MCT, monocarboxylate transporter; PDC, pyruvate dehydrogenase complex; PDK4, pyruvate dehydrogenase kinase 4; VLDL-TAG, very-low-density lipoprotein-triacylglycerol.

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diabetes (Saddik and Lopaschuk 1991). In addition, cardiac FA oxidation is very high upon reperfusion after ischemia (Kantor et al., 1999). Acetyl-CoA can be generated from a range of lipid sources: circulating non-esterified FA (NEFA); endogenous (stored) TAG; TAG in circulating very-low-density lipoproteins (VLDL); or circulating ketone bodies. In perfused rat hearts, a considerable rate of intramyocellular TAG breakdown supplies FA for post-ischemic fuel oxidation, and TAG lipolysis contributes about 20% of total FA oxidation even when hearts are perfused with exogenous FA (Saddik and Lopaschuk 1992).

3.6.2. Regulation of Cardiac Lipid Utilisation by PPAR The importance of PPAR in the adult heart appears relatively limited when the dietary or endogenous lipid supply is restricted, but PPAR assumes an important role in cardiac lipid management when lipid delivery to the heart is increased e.g., by fasting or by increasing dietary fat. In these situations, PPAR activation enhances the expression of genes encoding many proteins involved in FA transport, FA activation and FA oxidation (Figure 3.2): examples of PPAR-activated genes include muscle carnitine palmitoyltransferase 1 (m-CPT1) (which is crucial for the transport of long-chain FA into the mitochondria) and the -oxidation enzymes medium-chain acyl-CoA dehydrogenase (MCAD) and long-chain acyl-CoA dehydrogenase (LCAD). Increased expression of these genes allows increased mitochondrial FA oxidation. Thus, PPAR deficient (null) mice provided with free access to a high-carbohydrate low-fat diet do not show any obvious cardiac abnormalities, but starvation or high-fat feeding results in significant perturbations in cardiac lipid metabolism as a consequence of a failure of the heart to induce FA -oxidation pathways (Djouadi et al. 1998; Kersten et al. 1999; Leone et al. 1999), including abnormal accumulation of neutral lipid (Djouadi et al. 1998; Leone et al. 1999). Absence of PPAR decreases mitochondrial FA by approximately 70% (Aoyama et al. 1998; Campbell et al. 2002; Watanabe et al. 2000) due to reduced constitutive and/or inducible expression of several FA catabolic enzymes (Campbell et al. 2002).

3.6.3. Regulation of Cardiac Pyruvate Handling by PPAR Although PPAR is closely involved in the control of lipid handling, it also exerts a direct and vital influence on glucose utilisation at the level of the pyruvate dehydrogenase complex (PDC). Regulation of PDC activity is important since PDC inactivation is crucial for glucose conservation when glucose is scarce, whereas adequate PDC activity is required to allow both ATP and FA production from glucose. The regulatory kinase pyruvate dehydrogenase kinase (PDK) 4 is an important negative regulator of glucose oxidation, functioning via posttranslational modification of PDC (Sugden and Holness 2003) (Figure 3.2). Phosphorylation of E1 of PDC by PDK4 (and others of the PDK family) causes inactivation, sparing glucose and favouring FA oxidation. High saturated fat

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feeding increases cardiac PDK4 expression and PPAR activation acts synergistically with high-fat feeding to enhance cardiac PDK4 expression (Holness et al. 2002b). This essentially forces cardiac FA oxidation, despite available carbohydrate, and thereby optimises clearance of incoming lipid, particularly as the activity of this specific PDK isoform is increased by the increasing NADH/NAD+ concentration ratios that accompany FA -oxidation (BowkerKinley et al. 1998) (reviewed in Sugden and Holness 2003). In starvation, enhanced cardiac expression of PDK4 blocks glucose oxidation and again forces FA oxidation to maintain the ATP supply for cardiac contraction (Sugden and Holness 2003), this time in the face of a diminished supply of glucose. Increased expression of cardiac PDK4 in starvation is again linked to PPAR signalling, as well as to cardiac FA supply (Holness et al. 2002b; Wu et al., 2001). The involvement of lipid signalling via PPAR in the regulation of cardiac PDK4 expression, as well as that of FA oxidation enzymes, extends the operation of the classical glucose-FA cycle to the gene level and the newly-emerging field of nutrigenomics (Gillies 2003; Mutch et al. 2005).

3.6.4. Regulation of Hepatic Nutrient Handling by PPAR The liver is capable of either oxidising, or storing, FA at high rates - although not simultaneously. This metabolic versatility underlies its central position in wholebody lipid handling during the absorptive phase and after fasting. Incoming FA, derived from the diet or adipocyte TAG, can be re-esterified within the liver to provide the TAG component of VLDL: this function can be increased when hepatic FA delivery is higher than that required to fuel hepatic ATP requirements. During starvation, FAs generated by adipocyte lipolysis are taken up by the liver and oxidised to form the water-soluble lipid-derived fuels, the ketone bodies (Figure 3.3). The ketone bodies are avidly used as metabolic fuels by skeletal muscle and brain in starvation, reducing their requirement for glucose as metabolic fuel. PPAR regulates genes encoding regulatory proteins enhancing each stage of hepatic FA utilisation: FA uptake into the hepatocyte across the plasma membrane, FA retention within the cell and its total or partial mitochondrial oxidation, leading to ATP production and ketone body formation respectively (Figure 3.3). It is essential that the capacity for hepatic FA oxidation be up-regulated when lipid delivery is increased by a diet high in fat and also in starvation, where there is increased generation of FA derived from the hydrolysis of adipose-tissue TAG. These roles are best illustrated by the abnormal hepatic responses to starvation seen in PPAR–deficient mice (Kersten et al. 1999; Leone et al. 1999; Sugden et al. 2002). These animals exhibit an impaired ability to up-regulate hepatic FA oxidation in response to a high-fat diet, despite increases in FA supply. Secondary to impaired FA oxidation, PPAR deficient mice become hypoglycemic when fasted in part due to inadequate generation of ATP to fuel hepatic gluconeogenesis (Figure 3.3). Hypoglycemia also arises out of the necessity of heart and oxidative skeletal muscle to continue to use

3. Peroxisome Proliferator-Activated Receptors ketone bodies

glycerol AQP9

glucose

FA plasma membrane

GLUT

glycerol

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glucose

glycerol kinase

FA

acyl-CoA synthetase

glycerol 3-P gluconeogenesis pyruvate ATP

ketone bodies

FA-CoA

electron transport chain l-CPT1 ketone bodies

NADH/FADH2 CPT2

ketogenesis acetyl-CoA

TCA cycle β-oxidation

FA-CoA

Figure 3.3. Regulation of hepatic glucose and lipid utilisation by PPAR. Sites regulated by PPAR are shown in bold. Abbreviations: AQP, aquaporin; GLUT, glucose transporter; CPT, carnitine palmitoyltransferase; FA, fatty acid.

glucose at high rates, rather than to be able to switch to FA or ketone bodies as alternative energy fuels. Fasting hypoketonemia in PPAR null mice reflects a failure to enhance expression of the key mitochondrial enzyme in ketogenesis, mitochondrial HMG-CoA synthase (Le May et al. 2000). In response to lipolytic stimulation (exercise and fasting), adipocytes release not only hydrolysed FA, but also glycerol into the circulation. Glycerol is used for hepatic gluconeogenesis, a process involving its conversion to glycerol 3-phosphate by hepatic glycerol kinase (Figure 3.4). The contribution of glycerol to glucose production depends on the nutritional state: it may vary from 5% postprandially in humans (Baba et al. 1995), and approx 50% in the postabsorptive state in rodents (Peroni et al. 1995) to 20% in humans (Baba et al. 1995; Jensen et al. 2001) and >90% rodents after prolonged fasting (Peroni et al. 1995). It has been proposed that PPAR governs hepatic glycerol metabolism: the expression of several genes involved in the hepatic metabolism of glycerol

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glycerol plasma membrane

glucose

AQP glycerol

FA

GLUT glucose

FA acyl-CoA synthetase

glycerol-P

FA-CoA

esterification

glycerogenesis

TAG lactate

glycerol

FA

Figure 3.4. Actions of PPAR on the adipocyte. Sites regulated by PPAR are shown in bold. Abbreviations: AQP, aquaporin; GLUT, glucose transporter; FA, fatty acid; LPL, lipoprotein lipase.

(e.g. glycerol kinase, cytosolic and mitochondrial glycerol 3-phosphate dehydrogenase) is enhanced by fasting in wild type mice but not PPAR null mice (Patsouris et al. 2004). Furthermore, administration of pharmacological PPAR agonists lowers plasma glycerol concentrations (Patsouris et al. 2004). Under conditions of PPAR stimulation by the non--oxidisable FA tetradecylthioacetic acid, the liver functions as a major drain for circulating FA (Madsen et al. 2002). Furthermore, the fibrates in current clinical use primarily act on the liver in humans; hence, the TAG-lowering action of fibrates and related PPAR-targeted drugs in patients with hyperlipidemia is ascribed to increased FA clearance via oxidation primarily, but not exclusively in liver. However, new compounds are currently being developed to target PPAR in tissues other than the liver. Treatment of obese hypertriglyceridemic monkeys with a selective PPAR agonist increases LPL activity in muscle, with a reciprocal decrease in adipose tissue. Thus PPAR agonists suppress the uptake of TAGs in adipose tissue, while increasing uptake in muscle for oxidation (Bodkin et al. 2003).

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Regulation of Lipid Storage by PPAR

The human PPAR gene contains alternative transcription start sites resulting in three splice variants that produce three mRNA species in man (PPAR1, 2 and 3) and two in mice (PPAR1 and 2) (Fajas et al. 1997; Fajas et al. 1998). These mRNAs yield two distinct protein products: PPAR1and 3 mRNA encode an identical protein, PPAR1, while PPAR2 encodes a protein containing an additional 30 amino acids, PPAR2. PPAR1 is expressed in many tissues; PPAR2 is expressed predominantly in adipose tissue (Auboeuf et al. 1997). Conditioned PPAR knockouts have confirmed an essential role of PPAR in adipocyte differentiation and survival (Kubota et al. 1999; Rosen et al. 1999). Homozygous PPAR knockout mice die at embryonic day 10 from impaired placental development (Barak et al. 1999), highlighting that PPAR is also vital for nutrient and oxygen transfer to the fetus. PPAR interacts with a number of ligand-dependent co-activator complexes, including the CBP/p300 complex (Gelman et al. 1999), steroid receptor c-activator (SRC1) (Zhu et al, 1996), and PPAR coactivators PGC1 and PGC2, resulting in a range of biological activities. The ability of adipocytes to store FA as TAG in the fed state provides animals with a fuel store for use in time of need. In the fed state, insulin increases adipocyte TAG storage by augmenting adipocyte glucose uptake (allowing the production of glycerol 3-phosphate for FA esterification to form TAG), by direct stimulation of enzymes involved in the esterification of incoming FA, mainly generated locally via adipocyte LPL, and by effects to block TAG lipolysis. Adipocyte production of glycerol 3-phosphate is enhanced by PPAR activation (Guan et al. 2002; Picard and Auwerx 2002), which stimulates glycerol transport and glyceroneogenesis (Guan et al. 2002; Tordjman et al. 2003), in addition to glucose transport and LPL expression (Figure 4.4). Maintenance of glucose homeostasis requires optimisation of the body fat mass and distribution, as shown by animal models of obesity and lipodystrophy which are invariably insulin resistant (see e.g., Koutnikova et al. 2003). The use of PPAR knockout mice with tissue or isoform specific deletions, together with the identification of human subjects with mutations in PPAR, has highlighted its role as a critical regulator of adipose tissue differentiation and function, maintenance of fuel homeostasis and energy balance (Sewter and Vidal-Puig 2002). Mice harbouring dominant negative mutations of PPAR show an altered adipose-tissue distribution (Tsai et al. 2004). As ablation or impaired function of PPAR results in an insulin-resistant, lipodystrophic phenotype, the concept has been developed that the insulin-sensitising effects of PPAR activation (e.g., by the TZDs) reside in its actions in adipose tissue: these include, in addition to the induction of genes within mature adipocytes that allow them to take up and esterify FA to storage TAG, the recruitment of new adipocytes allowing an increase in lipid storage capability. Abnormal accumulation of TAG and other lipid derivatives in liver and muscle can result in insulin resistance (Shulman 2000). Thus, the sequestering of lipid within the adipocyte reduces the burden of excessive FA delivery to heart, skeletal muscle, and liver, where

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its oxidation and/or accumulation of FA-derived lipids could cause impaired glucose utilisation via substrate competition and/or insulin desensitisation (Frayn 2002). Of interest, specific deletion of the PPAR gene in fat and muscle causes insulin resistance (He et al. 2003; Hevener et al. 2003), but heterozygous PPAR knockout mice have improved insulin sensitivity, and are not susceptible to insulin resistance arising from a high-fat diet (Kubota et al. 1999). Thus partial activation is required to promote normal, but not excessive, adipose tissue stores. PPAR activation in the mature adipocyte also alters the expression and secretion of adipocyte proteins that regulate insulin sensitivity, resulting in an insulin-sensitising adipokine profile (Rangwala and Lazar 2004). Biologicallyactive adipokines include leptin (insulin sensitising) and TNF (which causes insulin resistance). Leptin is considered to be insulin sensitising through its effects to promote FA oxidation (Ceddia 2005). Increased leptin expression in PPAR+/− mice has been suggested to ameliorate insulin resistance, at least in part, through its effects to promote FA oxidation in liver and muscle (Yamauchi et al., 2001). This occurs in some tissues via activation of AMPK (Minokoshi et al. 2002). In addition, leptin treatment rapidly activates PPARs in C2C12 muscle cells (Bendinelli et al. 2005). TNF has an established role in the aetiology of insulin resistance (Hotamisligil 1999). PPAR suppresses the production of both leptin and TNF, the former allowing the adipocyte to continue to accumulate lipid and the latter relieving the insulin resistance that exposure to high levels of TNF induces. Recent emphasis has been placed on the insulin-sensitising adipokine adiponectin. Adiponectin appears to mediate its effects through two distinct receptors, AdipoR1 and AdipoR2: AdipoR1 is ubiquitous (highest in skeletal muscle), while AdipoR2 expression is more restricted (most abundant in liver) (Yamauchi et al. 2003). Primary hepatocytes bind full length adiponectin which stimulates PPAR ligand activity and AMPK activation (Yamauchi et al. 2003). The insulin-sensitising effects of the TZDs are impaired in mice lacking adiponectin (Kadowaki and Yamauchi 2005). Although PPAR is primarily involved in the regulation of lipid storage, it may also function in regulating glycerol transport across the adipocyte plasma membrane (Figure 3.4). The aquaporins are a family of small (approx 30kDa/monomer), integral membrane proteins. Aquaporin (AQP) 7 (also termed AQPap/7 in man), which transports glycerol as well as water, is expressed at high levels in adipose tissue (Kishida et al., 2000). During differentiation, 3T3-L1 adipocytes increase adrenaline-stimulated release of glycerol in parallel with the induction of AQP7 mRNA (Kishida et al. 2000). Knockdown of AQP7 in 3T3-L1 adipocytes by introducing small interfering mRNA suppresses adrenaline-stimulated glycerol release by approximately 50% (Maeda et al, 2004). AQP7 is a direct PPAR target gene in adipocytes (Guan et al. 2002; Kishida et al. 2001). Differentiated mouse and human adipocytes respond to the TZDs (ciglitazone and rosiglitazone) in culture with increased adipocyte AQP7 mRNA expression (Patsouris et al. 2004). The PPRE site in the human AQP7 promoter is important for high AQP7 mRNA expression in differentiated adipose cells and is responsible for the induction of AQP7 transcription by TZDs (Kondo et al. 2002). Contrasting with liver, adipose tissue normally contains

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very little glycerol kinase and glycerol liberated from adipocyte TAG is not recycled intracellularly to any great extent. However, it has been suggested (Guan et al. 2002), although it remains controversial (Tan et al. 2003a), that in concert with its effects to enhance adipose tissue AQP7 expression during adipocyte maturation, PPAR activation induces adipocyte glycerol kinase mRNA and activity, allowing glycerol to serve as a direct source of glycerol 3-phosphate within the adipocyte. Under physiological conditions, this could be potentially important for expansion of the adipose-tissue mass e.g., during puberty in females to promote the secretion of biologically active adipokines such as leptin, which are essential for reproductive function (Chehab et al. 1996; Chehab et al. 1997; reviewed by Chehab 1997). Interactions between PPAR and PPAR in the control of tissue development are not restricted to adipose tissue. The PPAR specific ligand WY14643 and the PPAR specific ligand GW0742 both induce increased expression of enzymes involved in mitochondrial FA -oxidation in cell cultures of neonatal rat cardiomyocytes (Cheng et al. 2004). The highest increment in PPAR and PPAR mRNA expression in the rat heart occurs from postnatal day 0 to day 7, whereas mRNA expression of MCAD and LCAD was observed between days 7 and 21 (Steinmetz et al. 2005). Interestingly, expression of PDK4, a PPAR/ PPAR target gene in the heart, also increases over this period (Sugden et al. 2000). Although the delay in the rise in MCAD and LCAD expression may reflect the absence of a necessary coactivator, the mRNA expression of the RXR receptors, PGC-1 and  do not parallel the rise in MCAD and LCAD. Other PPAR target genes in muscle include genes involved in the uptake and metabolism of glucose (e.g. c-Cbl associated protein and GLUT4) (Baumann et al. 2000).

3.8. The Role of PPAR in the Regulation of Oxidative Metabolism Relatively little was known about the function of PPAR until relatively recently, when the availability of PPAR-specific agonists and animal models allowed definition of the metabolic phenotype. PPAR is expressed early during embryogenesis and ubiquitously in adulthood (Braissant and Wahli 1998). Most PPAR-deficient mice die early in embryonic development due to a placental defect; those that do survive are characterised by reduced body fat mass, skin defects and altered myelinisation (Peters et al. 2000). PPAR is implicated in the regulation of proliferation of preadipocytes (Grimaldi 2001), keratinocytes (Tan et al. 2003b), enterocytes (Poirier et al. 2001) and placenta (Lim et al. 1999). Early studies also suggested that PPAR is involved in cholesterol metabolism (Oliver, Jr. et al. 2001) and adiposity (Barak et al. 2002). PPAR and PPAR are similarly abundant in heart (Muoio et al., 2002); it appears that in the heart many processes are dependent on functional PPAR (Muoio et al. 2002). Nevertheless, exposure of cardiac myocytes to either PPAR- or PPAR-specific

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agonists leads to significant induction of PPAR target genes involved in FA uptake and oxidation and both PPARs can be activated by FAs within this system. Skeletal muscle expresses all three PPAR isoforms, with high levels of PPAR expressed in both human and rodent skeletal muscle (Escher et al. 2001; Jones et al. 1995; Muoio et al. 2002). In this tissue, PPAR promotes oxidative metabolism and enhances aerobic capacity. Slow-twitch oxidative skeletal muscle is a major site of FA catabolism, particularly during starvation and long-term exercise, when circulating lipid delivery increases. However, most evidence suggests that signalling via PPAR impacts predominantly on the regulation of lipid gene expression and fat oxidation in fast-twitch, not slow-twitch, skeletal muscle, even though fast-twitch muscle does not normally oxidise FA as avidly as slow oxidative muscle (Dulloo et al. 2001; Holness et al. 2002a), and PPAR is predominantly expressed in mitochondrial-rich slow oxidative muscle (reviewed by Smith and Muscat 2005). Whereas PPAR ablation in the mouse results in abnormally high accumulation of neutral lipid in heart and liver in response to interventions increasing FA delivery, such as starvation, starvation of PPAR deficient mice leads to only minor abnormalities of skeletal-muscle FA metabolism (Muoio et al. 2002). In addition, skeletal muscles from PPAR null mice exhibit only minor changes in FA homeostasis and neither constitutive nor inducible mRNA expression of known PPAR target genes, including PDK 4 (Holness et al. 2002a) and uncoupling protein (UCP) 3 are negatively affected by the absence of PPAR (Muoio et al. 2002). PPAR may therefore assume greater importance in the modulation of lipid metabolism than PPAR in skeletal muscle, a conclusion supported by findings that PPAR mRNA expression levels in muscle are dramatically increased on fasting, under conditions where FA oxidation by skeletal muscle is increased. The failure of PPAR null mice to exhibit severe defects in skeletal muscle FA oxidation in response to starvation and exercise may reflect the fact that activation of PPAR using GW742 also increases FA oxidation and the mRNA levels of several classical PPAR target genes, including PDK4, m-CPT1, MCD and UCP3 (Muoio et al. 2002). Similarly, PPAR activation induces mRNA expression of UCP2 and UCP3 in human and L6 myotubes. This has led to the suggestion that significant overlap exists in the functions of PPAR and PPAR in mediating lipid-induced regulation of -oxidation pathways in skeletal muscle (Muoio et al. 2002). Gain- and loss-of-function experiments demonstrated that PPAR regulated the expression of genes encoding proteins involved in FA uptake, handling and -oxidation in differentiated mouse C2C12 myotubes (Holst et al. 2003). Similar observations were obtained using Affymetrix microarrays of rat L6 myotubes (Tanaka et al. 2003). Generation of a Cre/Lox transgenic mouse allowing skeletal muscle-specific overexpression of PPAR resulted in increased oxidative (fast-oxidative 2a) myofibers due to an increase in total myofiber number (Luquet et al. 2003). This response resulted in an increase in oxidative capabilities, characterised by up-regulation of succinate dehydrogenase, citrate synthase, heart-FA binding protein and UCP2. In addition, these mice exhibited net reduction of body fat mass related to reduced adipocyte size in fat depots.

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Subsequent studies demonstrated that overexpression of a constitutively-active PPAR in skeletal muscle resulted in protection against the development of diet-induced obesity (Wang et al. 2004). There is also evidence that PPAR is implicated in the response to physical exercise. The phenotypes elicited by muscle-specific overexpression of PPAR are similar to those resulting from endurance training in rodents (Allen et al. 2001) and humans (McCall et al. 1996) and overexpression of PPAR increases running endurance and resistance to fatigue (Wang et al. 2004). Repeated moderate exercise in mice also promotes up-regulation of PPAR in muscle (Luquet et al. 2003). Studies with synthetic PPAR ligands suggest that they may improve many factors associated with dyslipidemia, obesity and insulin resistance. Treatment of insulin-resistant obese monkeys with the potent PPAR agonist, GW1516, for 4 weeks normalises fasting plasma insulin and TAG concentrations, increases HDL-cholesterol and lowers LDL-cholesterol (Oliver, Jr. et al. 2001). Administration of GW1516 to obese mice reduced adiposity and improved insulin responsiveness (Tanaka et al. 2003; Wang et al, 2003). Mice expressing an activated mutant form of PPAR in brown and white adipocytes are characterised by up-regulation of genes involved in FA catabolism and energy uncoupling (Wang et al. 2003). These mice are also protected against diet-induced obesity (Wang et al, 2003). These studies suggest that therapeutic activation of PPAR could be used to decrease diet-induced obesity. Selective PPAR agonists (e.g. GW501516) are currently in clinical trials for the treatment of dyslipidemia, insulin resistance and obesity (Smith and Muscat 2005).

3.9. PGC-1: an Enhancer of Mitochondrial Function and Biogenesis Many metabolic programmes can be triggered by the expression of the transcriptional coactivator PGC-1. In addition to activating the PPARs, PGC-1 associates with other regulatory elements including cAMP response element (CREBP)-binding protein, the forkhead transcription factor FOXO1 and hepatocyte nuclear factor (HNF)-4, which have been implicated in the development of diabetes (Gupta et al. 2005; Nakae et al. 2002; Puigserver et al. 2003; Yamagata et al., 1996; Yoon et al. 2001). PGC-1 exhibits a tissue-enriched expression pattern (Knutti and Kralli 2001; Puigserver et al. 1998; Puigserver and Spiegelman 2003). PGC-1 is robustly expressed in a range of oxidative tissues, including heart and skeletal muscle, and is rapidly induced by a range of physiological conditions (cold exposure, acute exercise and fasting) that increase the demand for mitochondrial ATP or heat production (Baar et al. 2002; Goto et al. 2000; Irrcher et al. 2003; Lehman et al. 2000; Pilegaard et al. 2003; Puigserver et al. 1998; Terada et al. 2002; Terada and Tabata 2004; Wu et al. 1999). Its expression is also increased in the postnatal heart coincident with an increase in mitochondrial FA oxidation (Lehman et al. 2000).

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Recently, the PGC-1/ERR pathway has been demonstrated to regulate FA oxidation, oxidative phosphorylation and mitochondrial biogenesis in heart and skeletal muscle (Huss et al. 2004; Mootha et al. 2004; Schreiber et al. 2004; Willy et al. 2004). Evidence that PGC-1 regulates mitochondrial biogenesis in mammals includes studies of forced expression showing that PGC-1 induces expression of mitochondrial biogenic factors (e.g. nuclear respiratory factor 1 and 2 [NRF-1, NRF2] and the transcription factor of activated mitochondria [Tfam]) (Wu et al. 1999). In heart, PGC-1 induces mitochondrial biogenesis (Lehman et al. 2000) and, importantly, cooperates with PPAR to increase the expression of genes encoding mitochondrial FA oxidation enzymes, thereby increasing the capacity for FA oxidation and intracellular lipid clearance (Vega et al. 2000). In addition, forced expression of PGC-1 in skeletal muscle and heart triggers mitochondrial proliferation (Lehman et al. 2000; Lin et al. 2002; Russell et al. 2004). In parallel with the increase in PPAR signalling, fasting also elevates PGC-1 mRNA expression in liver (Herzig et al. 2001; Yoon et al. 2001). Importantly, PGC-1 mRNA expression is also increased in three different rodent models of increased hepatic gluconeogenesis (Herzig et al. 2001; Yoon et al. 2001). Increased expression of PGC-1 in liver via adenovirus vector enhances hepatic glucose production (Herzig et al. 2001) (reviewed in Vidal-Puig and O’Rahilly 2001). At least two clinical studies have identified a correlation between mutations of the Ppargc1a (previously known as the PGC-1 gene) and insulin resistance or diabetes (Ek et al. 2001; Hara et al. 2002). The effect of PGC-1 to induce gluconeogenesis appears to occur via an interaction with the FOXO1 pathway (Puigserver et al. 2003). In liver, FOXO1 and the glucocorticoid receptor, an additional PGC-1 partner, directly regulate PDK4 expression through consensus binding sites in the PDK4 gene promoter (Kwon et al. 2004). This effect both restricts glucose oxidation and also the provision of carbon for malonyl-CoA synthesis, thereby facilitating hepatic FA oxidation by relieving malonyl-CoA-induced inhibition of CPT 1 (Sugden and Holness 1994). Thus, while the biological role of PGC-1 to enhance glucose production in starvation is of major physiological importance, abnormally high expression/activity of this coactivator will also induce the phenotype of hepatic insulin resistance. PGC-1 expression is increased in heart after starvation (Lehman et al. 2000) and in skeletal muscle following exercise (Baar et al. 2002; Goto et al. 2000; Pilegaard et al. 2003). Skeletal muscle specific overexpression of PGC-1 results in a fibre-type switch from fast-twitch to slow-twitch fibres (Lin et al. 2002). The latter have an increased mitochondrial density compared with fasttwitch muscles, and thus an increased capacity for FA oxidation. Suppression of glucose utilisation (uptake/phosphorylation) makes a major contribution to glucose sparing during prolonged fasting (Holness and Sugden 1990) and, very recently, Kelly and colleagues have shown that PGC-1 directly activates PDK4 gene transcription in skeletal myotubes, leading to a reduction in glucose oxidation rates (Wende et al. 2005). This observation is important in view of our finding that signalling via PPAR is not obligatory for enhanced expression of PDK4 in skeletal muscle after starvation (Holness et al. 2002a).

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3.10. Dysregulation of PPARs and PGC-1 in Disease States Dysregulation of the PPAR axis and its control of cardiac FA utilisation have been detected in several cardiomyopathic disease states. Possibly as an adaptation to limited oxygen availability, in cardiac hypertrophy and heart failure the adult heart returns to a more “fetal” metabolic state, with greater reliance on glucose metabolism, (Calvani et al. 2000; van Bilsen et al. 2004). Oxygen is essential for ATP production from FAs, but glucose oxidation needs less oxygen per mole than FA oxidation and glucose utilisation can proceed, albeit less efficiently, in the absence of oxygen. However, less ATP is generated per mole of glucose oxidised, and when excess dietary lipid can be ingested, the potential exists for myocardial lipid accumulation and lipotoxicity. In animal models of heart failure related to pressure overload or ischemia, the expression of PPAR, PGC-1 and downstream target genes encoding FAO enzymes is diminished (Barger et al. 2000; Depre et al. 1998; Sack et al. 1996). Similar observations have been made in human patients with heart failure (Razeghi et al. 2002; Sack et al. 1996). It is implied that deactivation of the PPAR/PGC-1 axis is involved in the metabolic switch away from FA catabolism in the failing heart, with reversion to the fetal pattern of substrate use. Nevertheless, it remains controversial as to whether changes in cardiac PPAR signalling are adaptive or causally related to myocardial pathology. Although causality between the reciprocal changes in lipid and glucose utilisation has not been established, this metabolic switch may serve to preserve ventricular function within the context of pressure overload. The expression and activity of both PPAR and RXR are also suppressed in the hypoxic cardiac myocyte (Huss et al. 2001). Epidemiological studies reveal that people with diabetes mellitus are at high risk for the development of cardiovascular disease. The heart in uncontrolled diabetes is constrained from switching to glucose oxidation, and relies almost exclusively on lipid as an energy source (Belke et al. 2000; Lopaschuk 2002). In the diabetic heart the PPAR/PGC-1 axis is activated (Finck et al. 2002), suggesting that dysregulation of the PPAR/PGC-1 axis in diabetes is likely to be involved in the metabolic remodelling that occurs in diabetic heart to initially sustain the requirement for ATP. However, the precise role of these metabolic “switches” in the development of heart failure has not been clearly defined. Insight has been provided by supraphysiological activation of PPAR. Cardiacspecific over-expression of PPAR (myosin heavy chain [MHC- PPAR] mice), in the absence of alterations in lipid-fuel delivery, show increased expression of genes encoding FA oxidation enzymes, together with increased rates of myocardial lipid oxidation and decreased rates of glucose uptake and oxidation (Finck et al. 2002; Finck et al. 2003). This, however, is maladaptive, since these mice develop a cardiomyopathy with enhanced sensitivity to ischemic injury and recapitulate many of the metabolic abnormalities of the diabetic heart (Lopaschuk 1996; Rodrigues and McNeill 1992), including decreased expression of genes involved in cardiac glucose utilisation, accumulation of intracellular

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lipid, ventricular hypertrophy and systolic dysfunction. In this situation, the profile of myocardial TAG species is similar to the lipid species in the circulation, suggesting that the expanded lipid reservoir reflects increased uptake incompletely balanced by a corresponding increase in oxidation. When mice with cardiac-specific PPAR overexpression are made diabetic, they develop a more severe cardiomyopathy than wild-type controls (Finck et al. 2002). In contrast, PPAR null mice are protected from the development of a cardiomyopathy (Campbell et al. 2002). Effects of the TZDs to ameliorate hyperglycemia and hyperlipidemia in fatless mice demonstrate that, although predominantly expressed in adipose tissue, PPAR ligands may exert their effects via PPAR in tissues in addition to adipose tissue. Thus there is evidence that PPAR is expressed in non-adipose tissue under situations where other PPAR isoforms are functionally absent. Consistent with its function as an activator of lipogenesis and glycerolipid synthesis, elevated PPAR levels have been implicated as an early response in the progression of cardiac lipotoxicity (Muoio et al. 2002; Unger and Orci 2001). The PPAR agonist troglitazone prevents angiotensin II-induced hypertrophy in cultured rat cardiomyocytes (Asakawa et al. 2002) and similar results are obtained in endothelin 1-induced hypertrophy (Sakai et al. 2002). In humans with ischemic heart disease, an increase in PPAR expression has been observed when compared to patients suffering from dilated cardiomyopathy (Sakai et al. 2002). Although the underlying mechanism is not known, NF B may be involved in the signalling pathway (Takano et al. 2000; Zingarelli et al. 2003). The only cardiac gene that exhibits a clear change in expression in response to troglitazone treatment is acyl-CoA oxidase, which is involved in peroxisomal FA oxidation (Cabrero et al. 2003). However, care should be taken when interpreting these reports as the expression levels of PPARs may vary between the main cardiac cell types (cardiac myocytes, fibroblasts and endothelial cells) and the potential direct roles of PPAR in cardiac function are not yet clear (Kelly 2003). Of interest, the heart is closely associated with adipose tissue in adult man, and epicardial and pericardial depots can represent a significant fraction of the total mass of the heart, suggesting a possible role as a local FA source. With ageing and obesity, adipose tissue closely associated with the heart increases in mass. An intriguing possibility is that activation of PPAR in the cardiac-associated adipose tissue may act as a potential local sink for FA, sequestering excess FA when they are delivered in excess of the cardiac ATP requirement or when they cannot be cleared through oxidation. The heart expresses LPL and, interestingly, LPL activity in isolated cardiac myocytes can be inhibited by PPAR activation (Carroll and Severson 2001). This effect may potentially help protect against a potentially toxic oversupply of FAs to the myocardium. PPAR agonists have been reported to be cardioprotective against ischemic insult and to modify the cardiac hypertrophic growth response to pressure overload (Asakawa et al. 2002; Sivarajah et al. 2005). However, most cardiac effects are likely to be indirect because of the low level of expression of PPAR in the adult cardiac myocytes themselves, although the adipose tissue surrounding the heart itself may be targeted directly and act as a lipid buffer.

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Constitutive cardiac-specific overexpression of PGC-1 is lethal, the mice exhibiting high numbers of large mitochondria, developing a dilated cardiomyopathy (including depressed contractile function and massive edema) and ultimately dying by 6 weeks of age (Russell et al. 2005). As a consequence, subsequent studies employed the reverse tetracycline transactivator and a tetracycline responsive promoter to drive inducible cardiac-specific overexpression of PGC-1 (Russell et al. 2004). This model revealed developmental stagespecific responses to acute PGC-1 overexpression. Overexpression of PGC-1 during neonatal life causes dramatic proliferation of mitochondria, together with overt effects on cardiac function. Conversely, cardiac-specific overexpression of PGC-1 in adulthood resulted in only modest mitochondrial proliferation; however, it did elicit accumulation of aggregates of abnormal mitochondria (varied abnormal ultrastructure, including vacuoles, small granules and large membrane-bound inclusions within the mitochondria). In addition, cardiac dysfunction was observed in response to cardiac-specific overexpression of PGC1 in adulthood, including wall thinning and chamber dilation, together with the up-regulation of atrial natriuretic factor and brain natriuretic factor.

3.11.

Concluding Remarks

PPARs are central to whole-body and tissue specific lipid homeostasis, and by virtue of these actions, impact glucose homeostasis and insulin action. Activation of all three PPARs appears to decrease systemic lipid availability. PPAR and PPAR both promote mitochondrial FA oxidation, whereas PPAR acts to direct lipid towards storage in adipose tissue and away from oxidation in non-adipose tissues. Some physiological ligands for activation of each of the PPARs are PPAR subtype specific: such ligand specificity allows specificity of cellular downstream metabolic responses to each of the individual PPARs. The PPARs have proved to be attractive targets for the development of selective therapeutic agents for pharmacological correction of abnormal lipid metabolism in disease states. The newly-discovered role of the PPAR coactivator PGC-1 in both mitochondrial biogenesis and the coordination of lipid oxidation and glucose production provides a potential springboard for further research into mechanisms that may optimise muscle performance in exercise. Acknowledgements. The authors’ work cited in this review was supported by funding from the British Heart Foundation and Diabetes UK.

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4 Molecular Structure of the Mitochondrial Citrate Transport Protein Ronald S. Kaplan and June A. Mayor

4.1.

Introduction

4.1.1. Overview of the Mitochondrial Citrate Transport Protein The mitochondrial citrate transport protein (i.e., CTP) is located within the inner mitochondrial membrane and in higher eukaryotes catalyzes an obligatory 1:1 exchange of the dibasic form of a tricarboxylic acid (e.g., citrate, isocitrate, or cis-aconitate) for either another tricarboxylate, a dicarboxylate, or phosphoenolpyruvate (Palmieri et al. 1972; Bisaccia et al.1993; Palmieri 2004). The CTP occupies a prominent position within intermediary metabolism since following the efflux of citrate from the mitochondrial matrix and subsequent diffusion across the outer mitochondrial membrane, the resulting cytoplasmic citrate serves as the main carbon source fueling the fatty acid, triacylglycerol, and cholesterol biosynthetic pathways (Watson and Lowenstein 1970; Brunengraber and Lowenstein 1973; Endemann et al. 1982; Conover 1987). Additionally, via the sequential action of citrate lyase and malate dehydrogenase, cytoplasmic citrate enables the production of NAD+ , a cofactor that is required for glycolysis. Not only is CTP function essential to the bioenergetics of the physiological state, but altered functioning of this carrier in certain disease states such as diabetes (Kaplan et al. 1990b) and cancer (Kaplan et al. 1982) likely plays an important role in maintaining the aberrant bioenergetics that characterizes these pathologies.

The abbreviations used are: CTP, citrate transport protein; MTS, methanethiosulfonate; MTSES, sodium (2-sulfonatoethyl) methanethiosulfo-nate; MTSET, (2-(trimethylammonium)ethyl) methanethiosulfonate bromide; PAGE, polyacrylamide gel electrophoresis; S.E., standard error; and TMD, transmembrane domain. 97

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Due to the importance of the CTP, our laboratory has extensively studied the CTP’s structure/function relationships in order to understand its molecular mechanism. Thus the transporter has been purified in reconstitutively active form (Kaplan et al. 1990a), cloned (Kaplan et al. 1993), and overexpressed (Kaplan et al. 1995; Xu et al. 1995). Due to the high specific activity following reconstitution of the overexpressed yeast mitochondrial CTP (Kaplan et al. 1995), most of our recent investigative efforts have focused on this carrier source. Accordingly, we have constructed a Cys-less yeast CTP which retains wildtype functional properties (Xu et al. 2000) and have shown that in detergent micelles both the wild-type and the cys-less variants exist as homodimers (Kotaria et al. 1999). Each monomer is thought to contain six transmembrane domains (TMDs) (Kaplan et al. 1993; 1995). We then used the cysless CTP as a template upon which to reintroduce single cysteines at desired locations which were subsequently probed with biochemical and biophysical probes (Kaplan et al. 2000a; Kaplan et al. 2000b; Ma et al. 2004; Ma et al. 2005). Utilizing this approach we have identified the secondary structure, as well as the water-accessible and -inaccessible surfaces of TMDs III and IV. This data, in combination with examination of the ability of citrate (i.e., substrate) to protect against MTSES reagent mediated inhibition of CTP function (Ma et al. 2005) has enabled identification of portions of the CTP translocation pathway. Recently we have constructed a homology model of the CTP based on the crystal structure of the mitochondrial ADP/ATP carrier (Walters and Kaplan 2004). Superposition of our functional data onto the homology-modeled structure has permitted us to glean considerable additional insight into the CTP’s structurebased function. In this chapter, we will summarize our current state of knowledge regarding the structure/function relationships within the CTP (as of January 2006), and will delineate our assessment of where future efforts ought to focus.

4.1.2. Delineation of Essential Molecular Mechanistic Issues to Address in Understanding Transporter Function We begin with a comment regarding the nature of the issues that are important to resolve in order to attain a comprehensive understanding of the functioning of a metabolite transporter. These would include a complete definition of the substrate translocation pathway, the substrate binding site(s), the gates that control accessibility from one side of the membrane versus the other, and the detailed 3-dimensional conformational change(s) that occur during, and in fact represent the essence of, the transport process (West 1997). Portions of this information have been obtained with the CTP, whereas other aspects are currently under investigation. We now proceed with a detailed discussion of our current state of knowledge regarding the CTP molecular mechanism. For information regarding other aspects of this transporter the reader is referred to earlier reviews that are available (Kramer and Palmieri 1992; Kaplan and Mayor 1993; Kaplan 1996; Kaplan 2001).

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4.2. Identification of Residues that Comprise the Citrate Translocation Pathway 4.2.1. Preliminary Identification Based on MTS Reagent Accessibility We initiated identification of those residues that comprise the citrate translocation pathway via determination of the water-accessible surfaces of transmembrane -helices (Kaplan et al. 2000a; Ma et al. 2004). For these studies, starting with a Cys-less CTP, which displays native functional properties as the template (Xu et al. 2000), we engineered single cysteine residues at sequential positions within a given transmembrane domain. Following overexpression and functional reconstitution in liposomes of each single-Cys CTP variant, we examined the ability of two methanethiosulfonate (i.e., MTS) reagents to inhibit function. The ethylsulfonate and trimethylamine derivatives (i.e. MTSES and MTSET, respectively) are absolutely specific for cysteine, will not permeate a lipid bilayer in the absence of a translocation pathway, and react 5 × 109 more rapidly with the thiolate anion (that forms only when a cysteine sulfhydryl group is exposed to water) as compared to the unionized thiol group (Akabas et al. 1992; Holmgren et al. 1996; Karlin and Akabas 1998). Furthermore, as previously discussed, the molecular dimensions of these reagents are fairly similar to those of citrate (Xu et al. 2000), the latter also being a highly hydrophilic molecule. Thus, we believe that the MTS reagents and citrate will access similar aqueous domains within the CTP. Consequently, the theoretical underpinning for these studies consists of the idea that an accessible aqueous pathway through the lipid bilayer is likely to represent the substrate translocation pathway. This point will be further discussed below. Utilizing the above approach we have measured the accessibility of residues in TMDs III and IV to MTS reagents. It is noteworthy that, as depicted in Figures 4.1 (TMD IV) and 4.2 (TMD III) the rate constants for inactivation vary by greater than 7 and 5 orders of magnitude in TMDs IV and III, respectively. In certain cases, the rate constants for inactivation of adjacent engineered cysteines varied by 4–5 order of magnitude, thereby indicating a dramatic difference in accessibility to the aqueous environment. Moreover, the periodicity of the rate constant data strongly suggests that substantial portions of these TMDs exist in an -helical secondary structure. Furthermore, the rate constant data define a highly accessible surface of each helix, which likely comprises portions of the substrate translocation pathway, and a highly inaccessible surface of each helix which likely faces either the lipid bilayer or is tightly packed against other protein domains within the CTP. The results of computer modeling of these TMDs with the rate constant data superimposed on the models are depicted in Figure 4.2. As depicted in this Figure, each TMD displays a water-accessible face (depicted in red) and a water-inaccessible face depicted in blue. Moreover, TMDIII contains a series of moderately accessible residues depicted in magenta (see Figure 4.3, Panels C and D).

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4.2.2. Evaluation of Whether Citrate Protects Against the MTS-mediated Inhibition of CTP Function In order to test the concept that the water-accessible residues do in fact represent elements of the substrate translocation pathway through the CTP, we evaluated the ability of citrate to protect against the MTSES-mediated inhibition of CTP function (Ma et al. 2005). Thus, in the presence of a concentration of MTSES that yields 50–70% inhibition, we examined the ability of a spectrum of citrate concentrations to protect against the inhibition observed with 20 single-Cys TMDIII mutants. As depicted in Table 4.1, we observed protection at 8 sites with EC50 values that ranged 1.18–53.68 mM. Figure 4.4 depicts the TMDIII protectable residues in red and the nonprotectable residues in blue. Importantly, for the most part the protectable residues reside on one face of the -helix whereas the unprotectable residues reside on the opposite face. It is important to note that, with the exception of the S123C mutant, the other substrate-protectable residues display EC50 values that are significantly greater than their Km values. In general, the condition where substrate protection is observed with a given mutant, but where the measured EC50  Km value is diagnostic of a residue that lines the transport pathway, but is unlikely to have direct involvement in substrate binding (Fu et al. 2001; Ye and

 Figure 4.1. Rate constants for inactivation of citrate transport by MTS reagents versus location of engineered cysteine in TMD IV. Time course data for inactivation (activation) of a given single-Cys CTP variant by the MTS reagents were fitted to a simple exponential function by unweighted Marquardt nonlinear least squares: rt = r0 − r  • e−mct + r where rt is the observed activity remaining at time t (seconds), r0 is the initial activity, r is the asymptotic activity at t = , c is reagent concentration (M−1 ) and m is the rate constant for inactivation (M−1 sec−1 ). Estimates of m (which range over 7 orders of magnitude) are reliable as judged from replicate experiments producing standard errors less than 20% of each estimate. The rate constant for inactivation, m, can be interpreted as a measure of a residue’s accessibility to the MTS reagent. The broad range of estimates of m dictated that further analyses be confined to Log10 transformed values. For each data set (MTSES, MTSET and MTSEA) a Fourier analysis, as implemented in Mathematica, revealed a regular periodicity of 4 between residues 177 and 193 with a noticeable loss of signal when flanking sites were included. Therefore, further analyses were restricted to residues 177 through 193. For each MTS reagent, estimates of Log10 m were fitted to the periodic function Log10 m = sn − i + a • Sinn − x • 2/p using unweighted Marquardt nonlinear least squares. The magnitude of the rate constant for inactivation (Log10 m) is a function of position (residue number, n) with the straight line ((sn - i)) describing the slope (s) and intercept (i) of the overall trend in a data set and the trigonometric function a • Sinn − x • 2/p describing the periodicity either side of the line (p is the number of residues per 2 radians, a is amplitude and x a constant setting the register with respect to position, n). Reproduced from Kaplan et al., 2000a with permission from J. Biol. Chem.

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Figure 4.2. Rate constants for inactivation of citrate transport by MTS reagents versus location of engineered cysteine in TMD III. Time course data for inactivation of a given single-Cys CTP variant by either MTSES or MTSET were fitted to a simple exponential function by unweighted Marquardt nonlinear least squares as described in Figure 4.1. Estimates of m ranged over 5 orders of magnitude. The rate constant for inactivation, m, can be interpreted as a measure of a residue’s accessibility to the MTS reagent. The broad range of estimates of m dictated that further analyses be confined to Log10 transformed values.

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Figure 4.3. Computer modeling of residues 177–193 within TMDIV and 112–137 within TMDIII colored on the basis of reactivity toward the MTS reagents. TMD IV residues 177–193 (i.e., Panels A and B) were colored based on the mean of the deviations from the straight lines as depicted in Figure 4.1, Panel D. Residues with values less than zero are colored blue, while those with values greater than zero (i.e., more reactive and therefore more accessible) are colored red. The sequence was modeled initially as a standard -helix, with N- and C-terminal residues capped as amides. The structure was lightly minimized using the CHARMm force field (Brooks et al., 1983) to allow adjustment of the backbone around Pro177. Panel A: Space-filling representation, showing the face of the helix containing the more reactive residues (i.e., water-accessible). Panel B: Spacefilling representation, showing the face of the helix containing the less reactive residues (i.e., water-inaccessible). TMD III residues 112–137 (Panels C and D) were colored based on the mean MTS rate constant data depicted in Figure 4.2, Panel C. The most reactive, and thus most accessible residues (i.e., log10 rate constant values ranging from –2.72 –5.62 M −1 sec−1 ) are denoted in red. Residues of intermediate accessibility to the MTS reagents (i.e., log10 rate constant values ranging from –6.06 –6.31 M −1 sec−1 ) are denoted in magenta. Residues with the least accessibility to the MTS reagents (i.e., log10 rate constant values ranging from –6.91 –8.23 M −1 sec−1 ) are denoted in blue. Mutants for which MTS rate constant data could not be obtained (i.e., Cys substitution at positions 119 and 122) are denoted in gray. Panel C depicts a space-filling representation showing the face of the helix containing the more reactive (i.e., water-accessible) residues. Panel D depicts the face of the helix containing the less reactive (i.e., water-inaccessible) residues. Reproduced from Kaplan et al., 2000 and Ma et al., 2004 in modified form with permission from J. Biol. Chem. [See Color Plate I].

 Figure 4.2. For each data set (MTSES and MTSET) a Fourier analysis, as implemented in Mathematica, revealed a regular periodicity of 4 between residues 123 and 137 with a noticeable loss of signal when sites closer to the amino terminus were included. The gray traces indicate superimposed sinusoids. Panel A depicts the MTSES data. Panel B depicts results with MTSET. Panel C depicts the mean of the MTSES and MTSET data sets. Reproduced from Ma et al., 2004 with permission from J. Biol. Chem.

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Table 4.1. Kinetic Properties and EC50 Values for Those Single-Cys CTP Variants that Exhibited Substrate Protection Mutant

Vmax (nmol/min/mg)

Cys-less Gly-115 Leu-116 Gly-117 Leu-121 Ser-123 Val-127 Glu-131 Thr-135

1402 149 487 1448 640 104 229 13 267

± ± ± ± ± ± ± ± ±

69 5a 23a 4 36a 15a 29a 1a 32a

Km (mM) 0.471 0.388 0.228 0.355 0.300 2.502 0.136 0.170 0.118

± ± ± ± ± ± ± ± ±

0.052 0.005 0.008b 0.027 0.018 0.928a 0.006c 0.014c 0.005c

Vmax /Km (nmol/min/mg/mM)

EC50 (mM)

2977 (100%) 384 (12.9%) 2136 (71.7%) 4079 (137.0%) 2133 (71.7%) 42 (1.4%) 1684 (56.6%) 76 (2.6%) 2263 (76.0%)

– 4.15 1.18 2.78 5.31 6.66 1.84 7.18 53.68

a

Values are p < 0.01 from a two-tailed Student’s t test between the Cys-less CTP and individual single-Cys mutants, b values are p < 0.075, and c values are p < 0.05. Data from Ma et al., 2005 with permission from J. Biol. Chem.

Figure 4.4. Homology-modeled CTP TMD III colored according to substrate protection data. Red denotes those residues that were protected by citrate against inactivation by MTSES. Blue denotes residues with which no protection was observed. Panel A depicts the TMD III helical face containing residues protected by citrate facing outwards. Panel B depicts the face of the helix containing residues not protected by citrate. The helical view depicted in panel B is rotated approx. 180° relative to that depicted in panel A. Reproduced from Ma et al., 2005 with permission from J. Biol. Chem. [See Color Plate II].

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Maloney 2002). Furthermore, several other findings were unexpected and these will be discussed in the context of the homology modeled structure in sections 4.4.1–4.4.3.

4.2.3. Evaluation of the Temperature Dependence of Substrate Protection It is thought that two mechanisms can account for substrate protection against modification of a given residue by a covalent reagent (Seal and Amara 1998; Chen and Rudnick 2000; Leighton et al. 2002; Henry et al. 2003; Zomot and Kanner 2003; Gasol et al. 2004). One mechanism, involves occupancy of either the transport pathway or the substrate binding site by the substrate such that the latter can directly sterically impede the approach of a reagent to the modification site. The second mechanism involves a substrate-induced change in protein conformation that results in a decreased accessibility of a given residue to the modification reagent. Since the latter mechanism would presumably require substantive movement of protein domains, it would display a significant temperature dependence. In contrast, the first mechanism, which only involves collision and competition between substrate and the modification reagent, would display little temperature dependence. With these thoughts in mind, with the eight singleCys mutants with which substrate protection was observed, we examined the extent of this protection at 4°C versus 20°C (Ma et al. 2005). Similar levels of protection occurred at the two temperatures, thereby providing clear support for the concept that the citrate protection observed with these single-Cys CTP variants occurred by a direct steric blockage by substrate at the modification sites and thus citrate and MTSES both compete for sites that line the translocation pathway.

4.3. Construction of a Three-Dimensional Model of the CTP Through the efforts of my close collaborator and colleague D. Eric Walters homology modeling was used to construct a three-dimensional model of the CTP (Walters and Kaplan 2004) based on the x-ray crystal structure of the mitochondrial ADP/ATP transporter (Pebay-Peyroula et al. 2003). As depicted in Figure 4.5, superposition of the backbone trace of the homology-modeled CTP onto the crystallographically determined structure of the ADP/ATP carrier indicates that the CTP transmembrane domains are modeled quite well with a root mean square deviation of 0.94 Å, whereas there is more variability in the modeled loops with root mean square deviation values ranging from 0.72–2.06 Å. This structure has proven to be extremely valuable in that it enables the placement and interpretation of our functional data into a structural context, as well as in the design of fine-tuned testable hypotheses regarding the molecular functioning of the CTP (see sections 4.4, 4.6, and 4.7).

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B.

A. C2 C1

M3 M2 M1

Figure 4.5. Homology model of the CTP. Panel A: Superposition of the backbone trace of the best-scoring homology-modeled CTP (red) and the crystallographically determined ADP/ATP carrier (green). The intermembrane space loops (C1 and C2) and the mitochondrial matrix loops (M1, M2, and M3) connecting the transmembrane helices are labeled. Note the nearly identical positioning of the TMDs of the two carriers, which consequently appear as one color. Panel B: A ribbon diagram viewing the homology-modeled CTP from the outside (i.e., the intermembrane space). Panel A is reproduced from Walters and Kaplan, 2004 with permission from Biophys. J. [See Color Plate III].

4.4. Evaluation of CTP Functional Data in the Context of the Three-Dimensional CTP Homology Model Placing our functional data into a structural context has enabled the development of molecular explanations for several unexpected findings (Ma et al. 2005). These include the observed inability of citrate to protect against MTSES modification of cysteines substituted for either Leu120 or K134, despite the fact that both of these locations reside on the substrate-protectable face of TMDIII. In a similar vein, citrate protected against MTSES modification of the L121C mutant despite the fact that this position appears to reside on the non-protected surface of the TMDIII helix. The molecular explanations for these findings have proven to be of great interest and will now be discussed in detail.

4.4.1. Citrate Fails to Protect Against MTSES Modification of the Leu120Cys Mutant The explanation for this unexpected finding can be derived from close examination of the entire homology-modeled CTP structure (see Figure 4.6, Panel A). Thus it becomes apparent that TMDIII intersects TMDIV at an approximate

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Figure 4.6. Depiction of TMD III substrate protection data in the context of the CTP homology-modeled structure. Protected residues are shown in red, unprotected in blue, and Q182 in TMD IV in orange. For clarity, only the TMDs are depicted. Panel A: View into the transport pathway from the cytosolic surface of the bilayer. Panel B: Cross-sectional view of the transport pathway depicting TMDs II - IV. Helices I, V, and VI have been removed for clarity. This panel emphasizes the accessibility of L121 to the transport path. Reproduced from Ma et al., 2005 with permission from J. Biol. Chem. [See Color Plate III].

angle of 30°. When packed in this manner, the Leu-120 side-chain folds behind the side-chain of Gln-182 in TMDIV. Thus, we believe that the Gln-182 sterically blocks the approach of citrate to Leu-120, thereby preventing protection against MTSES modification. Importantly, as depicted in Figure 4.6 (Panel A), the Gln182 side chain is more proximal to the transport pathway, and as expected its modification by MTSES is protected by citrate (Ma et al. 2006).

4.4.2. Citrate Protects Against MTSES Modification of the Leu121Cys Mutant A second unexpected finding was the observation that even though within the TMDIII helix, Leu-121 is displaced approximately 100° from the substrate protectable surface of TMDIII, and thus would appear to reside on a portion of TMDIII pointing away from the transport path, we nonetheless observed substrate protection. However, as depicted in Figure 4.6 (Panel B), a cross-sectional view of the transport pathway clearly indicates the potential accessibility of this residue’s side-chain to the transport path. Additionally, the observation that both Leu-120 and Leu-121 reside near the intersection of TMDs III and IV, suggests that a low-magnitude helix-helix movement during the transport cycle may profoundly influence the accessibility of these residues to the transport pathway.

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4.4.3. Citrate Does Not Protect Against Modification of the Lys134Cys Mutant Based on the high MTS reagent accessibility of the Lys134Cys mutation (see Figure 4.2), and its location on the substrate-protectable surface of TMDIII, we were surprised to observe that citrate did not protect against MTSES-mediated inhibition. An examination of the homology-modeled TMDIII reveals that the Lys-134 side-chain resides near, but not entirely within, the transport path. We believe that our results support the idea that upon removal of the length and charge of the lysine side-chain, the resulting Cys side-chain is sufficiently distant from the transport path such that citrate does not access this sulfhydryl group with sufficient frequency to prevent covalent modification by MTSES. An important take-home message from these studies is that MTS reagent accessibility studies, when conducted in combination with substrate protection investigations, represent a powerful set of approaches enabling identification of residues that form the substrate translocation pathway. However, if only MTS reagent inhibition studies are conducted, then one must interpret the findings with appropriate caution. Thus, the MTS approach is, on its own, very effective for identifying water-accessible and -inaccessible surfaces of a TMD, as well as the likely secondary structure of such domains based on the periodicity of the accessibility data. However, when it comes to identification of those residues that line the translocation pathway one must interpret such data with caution realizing that there will likely be a small percentage of mistaken identification of such residues. Clearly, accessibility data in combination with substrate protection data is a much better indicator of translocation pathway lining residues.

4.4.4. Superposition of TMD III and IV Functional Data onto the CTP Homology-Modeled Structure Significant insight into CTP functioning is obtained upon superposition of our functional data onto the modeled CTP structure. For example, Figure 4.7 depicts the substrate-protectable residues of TMDIII and the MTS-accessible residues of TMDIV in red, and the residues in TMDIII that are not protected by substrate, as well as the MTS-inaccessible residues of TMDIV in blue. Finally, moderately accessible TMDIII residues are denoted in purple. This figure indicates that as one views the transport pathway from the external (i.e., cytosolic) surface of the bilayer, a substantial portion of the pathway is enclosed by TMD III and IV residues colored in red or purple. In contrast, the residues in these TMDs denoted in blue, face either other domains within the CTP or the lipid bilayer. Thus, our data clearly indicate the location of the pathway within the homology-modeled monomeric structure and offer firm support for the idea that two pathways exist per functional dimer. Also, as we have previously noted, Cys substitution mutagenesis with residues pointing into the pathway, is considerably more disruptive of CTP function than is mutagenesis of residues that do not comprise the pathway (Ma et al. 2004; Ma et al. 2006). Thus, of 14 essential

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Figure 4.7. Depiction of MTS rate constant data and substrate protection data superimposed onto the CTP homology model. Single-Cys CTP mutants with which the rate constants for MTS reagent inactivation have been determined are depicted in space-filling representation within the CTP homology-modeled structure. The substrate-protectable residues of TMDIII and the MTS-accessible residues of TMDIV are shown in red. Residues in TMDIII that are not protected by substrate, as well as the MTS-inaccessible residues of TMDIV are depicted in blue. Finally, moderately accessible TMDIII residues are denoted in purple. The view presented is looking down into the translocation pathway from the cytosolic surface of the bilayer. [See Color Plate IV].

residues that we have identified within the CTP, the side-chains of all but one project into the translocation pathway (Ma et al., 2006), implying that the precisely defined 3-dimensional architecture of the pathway is critical to support CTP function.

4.5. Criteria for Identification of Residues Involved in Substrate Binding Versus those Involved in Other Aspects of the Transport Mechanism An important issue regarding the molecular mechanism of any transport protein has to do with the identification of those residues that form the substrate binding site versus those residues that are important to other aspects of the translocation mechanism. We suggest that several criteria should be used in the identification

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of binding site residues. First, a residue that plays a prominent role in substrate binding should be essential for function, such that replacement of the residue via mutagenesis causes near total disruption of transporter function. The identification of an “essential” residue is relatively straightforward as it is obtained via a simple specific activity determination and typically involves residues, the mutation of which, causes a >98% inactivation of transporter function. This first condition represents a necessary but not sufficient criterion. Second, mutation of a substrate binding site residue should result in a profound increase in the Km value. Here we would expect that at least a 5-10-fold increase and often an even more profound increase would be obtained. Mutation of residues that are mechanistically essential but are not involved in substrate binding will often result in a large decrease in the Vmax with little change in the Km . Third, upon mutagenesis of a binding site residue to a Cys, we would expect substrate to protect against chemical modification of the engineered Cys, unless the remaining residues that comprise the substrate binding site are of sufficient distance from the engineered Cys such that citrate binding to these side-chains does not sterically block access of the MTS reagent to the cysteine sulfhydryl group. Fourth, the EC50 for protection should be similar to the Km value (Note: it is often overly rigorous to expect the EC50 to be exactly the same as the Km value, since an inhibitor that forms a covalent bond will ultimately overwhelm a substrate’s ability to competitively protect against this inhibition). Fifth, the substrate protection should be temperature-independent, thereby ruling out the possibility that protection arises indirectly as a consequence of a protein conformational change, but rather is due to a direct steric blockage by the substrate of the approach of the inhibitor to the site of modification. Sixth, if a non-transportable competitive inhibitor exists for the transporter, one would expect its Kd for binding to be greatly increased (>5–10-fold) upon mutation of a binding site residue to Cys. As mentioned above, with the CTP we have identified 14 essential residues (Ma et al. 2006). In a very recent development, we have been able to modify our transport assay such that we are now able to accurately measure Km values that are increased up to 100-fold over the native value. Current studies indicate that some of these essential residues display profound Km shifts. We believe that completion of these studies should enable an accurate, data-based modeling of the citrate binding site within the CTP during the next year.

4.6. The Location of the Monomer-Monomer Interface in Homodimeric Mitochondrial Transporters 4.6.1. Predictions With the CTP To date, extensive work from many laboratories utilizing a battery of approaches indicates that mitochondrial transporters exist in detergent micelles as homodimers and that this likely represents the functional form of these carriers (see Kotaria et al. 1999 for a review of this literature). For example with the CTP,

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utilizing both a gel filtration approach and a charge-shift native PAGE (polyacrylamide gel electrophoresis) approach, we observed that both the wild-type and the Cys-less CTP exist as homodimers (Kotaria et al. 1999). The question arises as to the role that a dimeric structure plays in the CTP mechanism. Insight into this issue can be gleaned from kinetic data indicating that the CTP operates by a sequential type mechanism which includes the formation of a ternary complex of two substrate molecules with the transporter prior to the translocation step (Bisaccia et al. 1993). This kinetic mechanism has important structural consequences in that two substrate binding sites must exist on opposite sides of the membrane and these sites must be exposed and occupied prior to the translocation event (Kramer and Palmieri 1992). Since superposition of our functional data onto the CTP homology-modeled structure indicates that each monomer contains a translocation pathway (Figure 4.7), a working model posits that the obligatory exchange catalyzed by the CTP occurs when a binding site within each monomer is occupied by substrate at opposite sides of the membrane. The two monomers in the homodimer must be out-of-phase (Pebay-Peyroula et al. 2003) and must closely communicate such that a conformational change leading to transport in one direction through one monomer is linked to a conformational change leading to transport in the opposite direction in the other monomer. Thus the composition of the dimer interface is essential to the correct functioning of the homodimer. We have proposed that TMDIII forms a substantive portion of the dimer interface (Ma et al. 2004; Walters and Kaplan 2004). Specifically we posit that G115 and G119, two residues which are highly conserved and which upon mutation to Cys considerably inactivate CTP function, form part of the interface and facilitate close packing between helices TMDIII and TMDIII’ (i.e., TMDIII in the other monomer). In addition, we have inferred that Ala118 also resides at the dimer interface. This supposition was based on our findings that the Ala118Cys CTP variant displayed a 7-fold increase in the Km , moderate accessibility to MTSES, and no substrate protection against MTSES-mediated inhibition of CTP function (Ma et al. 2004; Ma et al. 2005). These findings, in combination with our molecular modeling efforts, led to the conclusion that Ala118 may reside in close proximity to Ala118’ of the other monomer and together form an essential portion of the dimer interface. Further we posit that disruption of the interface by replacement with the bulkier Cys results in a substantial conformational change in the substrate binding site, leading to an increase in the observed Km . Interestingly, the Vmax value of the mutant also increases suggesting that Cys substitution at this site removes an intrinsic molecular constraint upon the transport rate, possibly one that is involved in the coordination of the function of the two monomers. A final hypothesis we have put forward regarding the composition of the dimer interface is depicted in Figure 4.8, and posits that E122 (a residue which upon mutation to Cys completely inactivates CTP function and which projects approximately 180° away from the transport pathway) of one monomer resides in close proximity to the TMDIII-TMDIV interface of the other monomer (Walters and Kaplan 2004). It is interesting to note, that the amide side-chain

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Figure 4.8. Schematic representation of a proposed dimer interface involving E122 of H3 and Q182’ of H4’. Helices 3 and 4 (i.e., H3 and H4) are identical to TMDs III and IV. Reproduced from Walters and Kaplan, 2004 with permission from Biophys. J.

of Q182’ (i.e., Q182 of the other monomer) resides at the TMDIII-TMDIV interface such that it can either face into the transport pathway or away from the pathway into the interface. Moreover, we have proposed that E122-Q182’ and E122’-Q182 interactions might serve to couple the conformational changes of the two translocation pathways, enabling the CTP to function as an obligatory antiporter.

4.6.2. Comparisons with the ADP/ATP Carrier Dimer Interface It should be noted that the above predictions regarding the dimer interface were developed by interpreting our functional data in the context of the CTP homology-modeled structure. An interesting question to consider is how these predictions compare with data available from the crystal structure of the mitochondrial ADP/ATP carrier. In the first structure, published by PebayPeyroula et al. (2003), only the monomeric form of the carrier was crystallized. However, in more recent studies crystals of two different dimeric forms of the transporter, which differ greatly in their orientation, were obtained (Nury et al. 2005). One of the crystal structures is compatible with existing data on the ADP/ATP carrier putative dimer interface and indicates the presence of cardiolipin at the interface. While the details of the protein-cardiolipin-protein contacts that they propose are beyond the scope of this chapter, in general they suggest that the interface involves interactions between cardiolipin, the matrix loops and TMDs II, IV, and VI, as well as possible interactions between the loops themselves. Presently, it is difficult to compare their findings with predictions that we have put forward for the CTP, since to date most of our work with the

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CTP has involved the TMDs and not the extramembranous loops. Accordingly, experiments are underway in our laboratory which are aimed at testing the role of the external loops in CTP function.

4.7.

Perspectives and Future Directions

We believe it accurate to say that substantial progress has been made in our understanding of the functioning of the CTP at the molecular level during the last 5 years, and yet in many ways we are now poised to address the most interesting questions. Thus, we believe we are close to attaining a detailed delineation of the citrate binding site(s). The precise detail of this site will be established via molecular modeling that will be guided by our extensive array of functional data. A second question that we are beginning to address has to do with the composition of the dimer interface. Our homology-model has permitted the development of a number of hypotheses concerning this matter, which we plan to test using a combination of functional assay and measurement of the distance between engineered cysteines utilizing both disulfide crosslinking and EPR. A third unresolved issue has to do with the conformational changes that occur during the opening and closing of the gates that modulate movement through the permeation pathway during the transport cycle. This can probably be best addressed via X-ray crystallographic analysis of trapped conformations representing different steps of the transport cycle. Towards this end, we are in the midst of extensive crystallization trials utilizing Fab fragments generated from conformation-specific anti-CTP monoclonal antibodies, various CTP inhibitors, and inactive CTP mutants that may be locked in a given conformation. Through a combination of the above approaches we believe that we will achieve a complete understanding of the molecular basis for CTP functioning during the next 5–10 years. Acknowledgements. The authors would like to acknowledge the efforts of a talented list of colleagues whom we have had the good fortune of working with and who have made substantial contributions towards conducting much of the work described in this chapter. This list includes Chunlong Ma, Sreevidya Remani, as well as Drs. Rusudan Kotaria, Yan Xu, and D. Eric Walters. This work was supported by National Institutes of Health Grant GM-054642 to R.S.K.

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Brunengraber H, Lowenstein JM (1973) Effect of (-)-hydroxycitrate on ethanol metabolism. FEBS Lett 36:130–132 Chen J-G, Rudnick G (2000) Permeation and gating residues in serotonin transporter. Proc Nat Acad Sci. USA 97: 1044–1049 Conover TE (1987) Does citrate transport supply both acetyl groups and NADPH for cytoplasmic fatty acid synthesis? Trends Biochem Sci. 12:88–89 Endemann G, Goetz PG, Edmond J, Brunengraber H (1982) Lipogenesis from ketone bodies in the isolated perfused rat liver. Evidence for the cytosolic activation of acetoacetate. J Biol Chem 257:3434–3440 Fu D, Sarker RI, Abe K, Bolton E, Maloney PC (2001) Structure/function relationships in OxlT, the oxalate-formate transporter of oxalobacter formigenes. Assignment of transmembrane helix 11 to the translocation pathway. J Biol Chem 276: 8753–8760 Gasol E, Jimenez-Vidal M, Chillaron J, Zorzano A, Palacin M (2004) Membrane topology of system xc-light subunit reveals a reentrant loop with substrate-restricted accessibility. J Biol Chem 279:31228–31236 Henry LK, Adkins EM, Han Q, Blakely RD (2003) Serotonin and cocaine-sensitive inactivation of human serotonin transporters by methanethiosulfonates targeted to transmembrane domain I. J Biol Chem 278: 37052–37063 Holmgren M, Liu Y, Xu Y, Yellen G (1996) On the use of thiol-modifying agents to determine channel topology. Neuropharmacology 35:797–804 Kaplan RS (1996) Mitochondrial transport processes. In: Schultz SG, Andreoli T, Brown A, Fambrough D, Hoffman J, Welsh J (eds) Molecular Biology of Membrane Transport Disorders. Plenum Press, New York, pp. 277–302 Kaplan RS (2001) Structure and function of mitochondrial anion transport proteins. J Membrane Biol 179:165–183 Kaplan RS, Morris HP, Coleman PS (1982) Kinetic characteristics of citrate influx and efflux with mitochondria from Morris hepatomas 3924A and 16. Cancer Res 42:4399– 4407 Kaplan RS, Mayor JA, Johnston N, Oliveira DL (1990a) Purification and characterization of the reconstitutively active tricarboxylate transporter from rat liver mitochondria. J Biol Chem 265:13379-13385 Kaplan RS, Oliveira DL, Wilson GL (1990b) Streptozotocin-induced alterations in the levels of functional mitochondrial anion transport proteins. Arch Biochem Biophys 280:181-191 Kaplan RS, Mayor JA, Wood DO (1993) The mitochondrial tricarboxylate transport protein. cDNA cloning, primary structure, and comparison with other mitochondrial transport proteins. J Biol Chem 268:13682-13690 Kaplan RS, Mayor JA, Gremse DA, Wood DO (1995). High level expression and characterization of the mitochondrial citrate transport protein from the yeast Saccharomyces cerevisiae. J Biol Chem 270:4108–4114 Kaplan RS, Mayor JA, Brauer D, Kotaria R, Walters DE, Dean AM (2000a) The yeast mitochondrial citrate transport protein: probing the secondary structure of transmembrane domain IV and identification of residues that likely comprise a portion of the citrate translocation pathway. J Biol Chem 275:12009–12016 Kaplan RS, Mayor JA, Kotaria R, Walters DE, Mchaourab HS (2000b) The yeast mitochondrial citrate transport protein: Determination of secondary structure and solvent accessibility of transmembrane domain IV using site-directed spin labeling. Biochemistry 39:9157–9163

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Karlin A, Akabas MH (1998) Substituted-cysteine accessibility method. Methods Enzymol 293:123–145 Kotaria R, Mayor JA, Walters DE, Kaplan RS (1999) Oligomeric state of wild-type and cysteine-less yeast mitochondrial citrate transport proteins. J Bioenerg Biomemb 31:543–549 Kramer R, Palmieri F (1992) Metabolite carriers in mitochondria. In: Ernster L (ed) Molecular Mechanisms in Bioenergetics. Elsevier, New York, pp. 359–384 Leighton BH, Seal RP, Shimamoto K, Amara SG (2002) A hydrophobic domain in glutamate transporters forms an extracellular helix associated with the permeation pathway for substrates. J Biol Chem 277: 29847–29855 Ma C, Kotaria R, Mayor JA, Eriks LR, Dean AM, Walters DE, Kaplan RS (2004) The mitochondrial citrate transport protein: probing the secondary structure of transmembrane domain III, identification of residues that likely comprise a portion of the citrate transport pathway, and development of a model for the putative TMDIII-TMDIII’ interface. J Biol Chem 279:1533–1540 Ma C, Kotaria R, Mayor JA, Remani S, Walters DE, Kaplan RS (2005) The yeast mitochondrial citrate transport protein: characterization of transmembrane domain III residue involvement in substrate translocation. J Biol Chem 280:2331–2340 Ma C, Remani S, Kotaria R, Mayor JA, Walters DE, Kaplan RS (2006) The mitochondrial citrate transport protein: evidence for a steric interaction between glutamine 182 and leucine 120 and its relationship to the substrate translocation pathway and identification of other mechanistically essential residues. Biochim Biophys Acta, Manuscript Submitted Nury H, Dahout-Gonzalez C, Treqeguet V, Lauguin G, Brandolin G, Pebay-Peyroula E (2005) Structural basis for lipid-mediated interactions between mitochondrial ADP/ATP carrier monomers. FEBS Lett 579:6031–6036 Palmieri F (2004) The mitochondrial transporter family (SLC25): physiological and pathological implications. Pflugers Arch - Eur J Physiol 447:689–709 Palmieri F, Stipani I, Quagliariello E, Klingenberg M (1972) Kinetic study of the tricarboxylate carrier in rat liver mitochondria. Eur J Biochem 26:587–594 Pebay-Peyroula E, Dahout-Gonzalez C, Kahn R, Trezeguet V, Lauquin GJ-M, Brandolin G (2003) Structure of mitochondrial ADP/ATP carrier in complex with carboxytractyloside. Nature 426:39–44 Seal RP, Amara SG (1998) A reentrant loop domain in the glutamate carrier EAAT1 participates in substrate binding and translocation. Neuron 21: 1487–1498 Walters DE, Kaplan RS (2004) Homology-modeled structure of the yeast mitochondrial citrate transport protein. Biophys J 87:907–911 Watson JA, Lowenstein JM (1970) Citrate and the conversion of carbohydrate into fat. Fatty acid synthesis by a combination of cytoplasm and mitochondria, J Biol Chem 245:5993–6002 West IC (1997) Ligand conduction and the gated-port mechanism of transmembrane transport. Biochim Biophys Acta 1331:213–234 Xu Y, Mayor JA, Gremse D, Wood DO, Kaplan RS (1995) High-Yield bacterial expression, purification, and functional reconstitution of the tricarboxylate transport protein from rat liver mitochondria. Biochem Biophys Res Comm 207: 783–789 Xu Y, Kakhniashvili DA, Gremse DA, Wood DO, Mayor JA, Walters DE, Kaplan RS (2000) The yeast mitochondrial citrate transport protein: probing the roles of cysteines, Arg181, and Arg189 in transporter function. J Biol Chem 275:7117–7124

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Ye L, Maloney PC (2002) Structure/function relationships in OxlT, the oxalate/formate antiporter of Oxalobacter formigenes: assignment of transmembrane helix 2 to the translocation pathway. J Biol Chem 277: 20372–20378 Zomot E, Kanner BI (2003) The interaction of the gamma-aminobutyric acid transporter GAT-1 with the neurotransmitter is selectively impaired by sulfhydryl modification of a conformationally sensitive cysteine residue engineered into extracellular loop IV. J Biol Chem 278: 42950–42958

5 Regulation of Pyruvate and Amino Acid Metabolism Thomas C. Vary, Wiley W. Souba and Christopher J. Lynch

5.1.

Introduction

Amino acids serve multiple functions in body ranging from the basic building blocks of proteins to precursors for glucose and urea synthesis. The circulating amino acids are generally categorized with respect to either their function (gluconeogenic or ketogenic) or availability (essential or nonessential). The metabolism of amino acids is directly controlled through mitochondrial function. In addition, there is growing evidence that amino acids participate in processes that control cellular functions. Amino acids are not only utilized as metabolic substrates, increasingly it has become evident they also participate in the regulation of various cellular processes including protein synthesis, glycogen synthesis and autophagy. The regulatory processes affected by amino acids may involve mitochondria. This chapter will provide an overview of the current understanding of the role of the mitochondrial function in controlling amino acid metabolism.

5.2. Metabolism of Amino Acids via Pyruvate Dehydrogenase The complexity of the processes involved in utilization and synthesis of glucose represent one of the fundamental metabolic processes of the human body. The overriding reason for the complexity of glucose homeostasis is the central nervous system’s requirement for glucose to sustain energy production. The liver and kidney represent the major organ systems capable of gluconeogenesis in man. Skeletal muscle, by virtue of its mass, represents a major source of gluconeogenic substrates. As such there is an intricate balance in the release of amino acids from protein stores in muscle and the utilization of amino acids for gluconeogenesis in the liver. Thus, interorgan transfer of carbon skeletons derived from amino acids contributes significantly to the flux of nutrients delivered to the brain. In the most simplistic scenario, when glucose becomes unavailable via ingestion of food, amino acid availability is increased by net protein degradation in skeletal muscle. There are several fates of the amino acids released by proteolysis in 117

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muscle (Figure 5.1). First, a certain amount of amino acids can be reincorporated into proteins via protein synthesis. Second, amino acids can undergo oxidation in mitochondria producing in the process energy. At least six amino acids (alanine, aspartate, glutamate, leucine, isoleucine and valine) can be oxidized by skeletal muscle. Third, amino acids can be released into the venous blood system. All amino acids are readily released from skeletal muscle via specialized transport systems. Fourth, skeletal muscle has the ability to interconvert the nitrogen from amino acids to alanine and glutamine, thereby promoting de novo synthesis of these two amino acids. Mitochondrial regulation of amino acid and carbohy-

Figure 5.1. Interrelationship between mitochondrial amino acid and pyruvate oxidation in skeletal muscle. LEU – leucine; BCATm – Mitochondrial branched chain amino transferase; -KG – -ketoglutarate; KIC – -ketoisocaproic acid; BCKD – branched chain amino acid dehydrogenase; TCA – tricarboxylic acid cycle; PDH – pyruvate dehydrogenase complex; PYR – pyruvate; ALA – alanine; GLUT – glutamate.

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drate oxidation plays a vital role in the extent of de novo alanine and glutamine synthesis. Indeed, the majority of amino acids released by skeletal muscle during starvation are alanine and glutamine (Felig 1975). The total content of alanine plus glutamine in skeletal muscle proteins is slightly more than 10%, whereas they comprise as much as 60% of amino acids released from muscle. Hence, these two amino acids are released in amounts greater than can be accounted for by proteolysis of skeletal muscle proteins. Hence, they undergo “de novo” synthesis (Figures 5.2 and 5.3).

Figure 5.2. Relationship between branched chain amino acid metabolism and glutamine synthesis in skeletal muscle. BCKD – branched chain amino acid dehydrogenase; TCA – tricarboxylic acid cycle.

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Figure 5.3. Mitochondrial regulation of interorgan amino acid fluxes between skeletal muscle and liver. LEU – leucine; BCATm – mitochondrial branched chain amino transferase; ; -KG – -ketoglutarate; KIC – -ketoisocaproic acid; BCKD – branched chain amino acid dehydrogenase; TCA – tricarboxylic acid cycle; PDH – pyruvate dehydrogenase complex; PYR – pyruvate; ALA – alanine; GLUT – glutamate; OAA – oxaloacetate; PEP – phosphoenolpyruvate; MITO – mitochondria.

Amino acids are released from the skeletal muscle and are taken up by liver and converted into glucose with the release of the nitrogen group being detoxified via the formation of urea (Figure 5.3). Urea is then excreted from the body via normal kidney function. The physiological significance of amino acid oxidation

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for energy is not thought to be as important as the ability of skeletal muscle to convert amino acids to alanine and glutamine that are released in quantities that greatly exceed their concentration in muscle proteins.

5.2.1. Alanine Cycle The pathway by which alanine is formed and transported back to liver has been referred to as the glucose/alanine cycle. About 30% of the alanine released by skeletal muscle is derived from net protein degradation, while the remaining 70% is synthesized de novo. Glucose carbon is metabolized in skeletal muscle to pyruvate via the glycolytic pathway. Once pyruvate is formed, there are essentially three pathways that it can follow in muscle tissues. First, it can be transported into the mitochondria, and oxidized via the pyruvate dehydrogenase complex to acetyl-CoA and then subsequently completely oxidized via the tricarboxylic acid cycle. Second, it can be reduced to lactate with the consumption of NADH. Lastly, pyruvate can be transaminated to alanine in a reaction catalyzed by the alanine aminotransaminase. The pyruvate dehydrogenase (PDC) complex catalyzes the first irreversible reaction in the mitochondrial oxidation of glucose carbon and as such is the primary determinant of glucose oxidation (Randle et al. 1984; Randle et al. 1988; Randle 1998; Sugden et al. 1989). The regulation of this complex is important to glucose homeostasis. Oxidation of pyruvate results in the depletion of body glucose carbon sources because glucose cannot be synthesized from acetyl-CoA and the PDH reaction is physiologically irreversible (Randle et al. 1984; Randle et al. 1988; Randle 1998; Sugden et al. 1989). Thus, the activity of the PDH complex determines whether pyruvate is oxidized via the Krebs cycle or converted to lactate via lactate dehydrogenase or converted to alanine via alanine transaminase. The source of the nitrogen for de novo alanine synthesis occurs via transaminations with other amino acids. As much as 60% of the nitrogen for de novo alanine synthesis is derived from branched chain amino acids (valine, isoleucine, and leucine) (Hayman and Miles 1982). For the branched chain amino acids, transamination is catalyzed by the branched chain aminotransferase. There are two aminotransferases; one that has a preference for valine (a deficiency of which gives rise to hypervalenemia) and one that has a preference for leucine or isoleucine (a deficiency of which gives rise to leucine-isoleucinaemia). The resulting -keto acids (-ketoisocaproic; -keto3-methylvalarate; -ketoisovalate) are available for oxidative decarboxylation via the mitochondrial branched chain amino acid dehydrogernase (BCKD) an enzyme located in the inner mitochondrial membrane (Figure 5.1) and eventually metabolized to either acetyl-CoA (leucine) or succinyl CoA (isoleucine and valine) which enter the citric acid cycle. A deficiency in BCKD leads to the accumulation of the -keto acids in the blood and subsequent elimination from the body (see section on Maple syrup urine disease).

5.2.2. Glutamine Cycle Glutamine serves various metabolic functions in different tissues and under different physiologic states. Its functions within the cell are characterized by its

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(a) utility in nitrogen transport and ammonia detoxification (b) importance in maintaining the cellular redox state (c) position as a metabolic intermediate and (d) and ability to serve as an energy source. Glutamine like alanine must arise from amino acid metabolism in skeletal muscle. Glutamine serves as the primary ammonia shuttle between tissues. This “nitrogen shuttle” provides a non-toxic means of transporting ammonia from the periphery to the viscera. After reaching its target organ ammonia can be regenerated for excretion or ureagenesis. Glutamine is synthesized from intramuscular glutamate and ammonia via glutamine synthetase (Figures 5.2 and 5.4). The glutamine synthetase (GS) pathway accounts for as much as 50% of the glutamine released with elevated rates of glutamine release (Vary et al. 1989a; Vary and Murphy 1989). Expression of GS is increased in muscle during a number of catabolic states, with induction of GS mRNA levels by 400–700% (Abcouwer et al. 1996; Abcouwer et al. 1997; Ardawi and Majzoub 1991; Labow et al. 2001). Induction of GS gene expression in skeletal muscle is largely dependent on adrenal derived hormones (Abcouwer et al. 1995; Chandrasekher et al. 1999; Lukaszewicz et al. 1997). Glutamate is most likely formed from 2-oxoglutarate via transamination from other amino acids. The 2-oxoglutarate arises from mitochondrial citrate and transport of the carbon out of the mitochondria via malate/aspartate shuttle. The ammonia arises from the reactions catalyzed by the glutamate dehydrogenase

Figure 5.4. Glutamine balance showing relationship between availability and consumption.

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reaction. Activation of PDH complex activity through dichloroacetate leads to increased glutamine production presumably by lowering the availability of pyruvate for transamination with glutamate to form alanine (Vary et al. 1988b). Hence, the supply of glutamate would be elevated and available for de novo glutamine synthesis. The liver serves as the major regulatory tissue responsible for whole-body ammonia detoxification and glutamine homeostasis (Haussinger et al. 1992; Meijer et al. 1990). Transport of glutamine across the plasma membrane of liver cells may represent a rate-limiting step in its metabolism, especially when intracellular catabolism is accelerated (Haussinger et al. 1985; Low et al. 1993). In normal rat and human hepatocytes, glutamine transport is predominantly mediated by a Na+ -dependent transporter with narrow substrate specificity (glutamine, histidine, and asparagine) termed system N (Bode et al. 1995; Kilberg et al. 1980). In the mitochondria, ammonia is derived from glutamine by hydrolysis of glutamine to glutamate by the enzyme glutaminase. Hydrolysis of glutamate to -ketoglutarate by the enzyme glutamate dehydrogenase can also generate ammonia. In the liver, ammonia can combine with CO2 to form carbamoyl phosphate that subsequently is detoxified by the urea cycle. The amino group on glutamate and glutamine may also enter the urea cycle directly through a reaction involving oxaloacetate to generate aspartate. One of the major functions of the liver is the maintenance of ammonia homeostasis within the body. Glutamine metabolism is distributed heterogeneously throughout the liver with glutamine consumption via glutaminase concentrated in the periportal hepatocytes while glutamine synthesis via glutamine synthetase is localized within the perivenous hepatocytes. This distribution provides the liver with an elegant mechanism to detoxify the blood of ammonia while contributing glutamine to the systemic supply. Maintenance of acid-base homeostasis is an essential function of the kidney, particularly under conditions associated with metabolic acidosis. During metabolic acidosis, aminogenesis is higher in kidney, allowing for the excretion of protons in the form of ammonium ions in urine. The use of glutamine as a nitrogen shuttle is also important in the excretion of nitrogenous wastes and the maintenance of acid-base homeostasis. Circulating glutamine contributes 80–90% of the ammonia produced during acidosis. Under normal conditions, renal uptake of glutamine from the blood is minimal. During chronic metabolic acidosis, however, renal extraction of glutamine increases markedly. Enhanced renal glutamine uptake and metabolism results from a coordinated series of events triggered by the disruption of normal acid-base balance. The homeostatic response involves alterations in interorgan glutamine flux, such that glutamine release from skeletal muscle is doubled, the splanchnic bed shifts from net glutamine uptake to glutamine release, and the kidney becomes the major site of glutamine consumption. Three Na+ -dependent amino acid transport systems (system A, system ASC (ATB0), and system N) transport glutamine across cell membranes. Almost all the glutamine filtered at the glomerulus is reclaimed in the convoluted proximal tubule, leaving little margin for increased glutamine uptake from the tubular

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lumen during acidosis. The major portion of the increased renal glutamine taken up enters the tubular epithelium across the basolateral membrane from the blood via system N1 (SN1) amino acid transporter. Acidosis causes a 10-fold increase in whole kidney SN1 mRNA level and a 100-fold increase in the cortex (Karinch et al. 2002). Acidosis increases Na+ -dependent glutamine uptake (SN1) into basolateral and brush-border membrane vesicles isolated from rat cortex. The SN1 carrier appears to be the most important cell membrane transporter of circulating glutamine into the renal tubular cell during acidosis. Once circulating glutamine is transported into the renal tubular cell, it is acted upon by a kidney-specific isozyme of glutaminase; hydrolyzing glutamine and generating ammonia for excretion in the urine (Deferrari et al. 1994; Dejong et al. 1993). The catabolism of glutamine provides the majority of nitrogen excreted in the urine. Furthermore, the availability of ammonia also facilitates the excretion of acid loads by conjugating H+ ions and generating ammonium ions for excretion. The metabolism of glutamine produces bicarbonate ions that can function to further neutralize hydrogen ions in the blood. The adaptive changes in proximal renal tubules include 1) escalating glutamine transport both in brush border and basolateral membranes increases the availability of glutamine for ammoniagenesis and simultaneously augments secretion of hydrogen ions into urine and blood, 2) not surprisingly, glutamine consumption via phosphatedependent glutaminase, glutamate dehydrogenase, and phosphoenol carboxykinase become accelerated within the kidney during acidosis (Deferrari et al. 1994; Deferrari et al. 1997), and 3) upsurges in activity of Na+ /H+ antiporter and Na+ /HCO3 − co-transporter increases H+ secretion into urine and bicarbonate ion reabsorption into the blood. Aside from ureagenesis, glutamine plays a key role in regulating glutathione synthesis. Glutathione is a tripeptide composed of glutamate, cysteine and glycine and represents the major source of cellular reducing equivalents, protecting the cell against oxidative injury. As a source of intracellular glutamate, glutamine provides one of the constituents of glutathione, but glutamine can also contribute to glutathione synthesis indirectly by exchanging intracellular glutamate (derived from glutamine) across the cell membrane for extracellular cysteine, another component of glutathione. The importance of glutathione resides in its ability to catalyze the reduction of organic hydroperoxides and hydrogen peroxide to limit reactive oxygen species damage. Glutathione peroxide lowers oxygen reactive oxygen species through several mechanisms. First, it converts hydrogen peroxide to water. Second, it converts peroxidated fatty acids to hydroxyl fatty acids. Hydroxyl radicals tend to interact with unsaturated fatty acids in phospholipids and other lipid components of membranes forming lipid hydroperoxides ultimately resulting in increased hydrophilicity of membranes and inhibition of several enzymes. The flow of glutamine into and out of the liver can vary widely. Physiological concentrations of ammonia within the portal circulation can produce a ‘feedforward stimulation’ of hepatic glutaminase (Haussinger 1990), allowing the liver to increase glutamine consumption when glutamine is abundant (Welbourne and Oshi 1990). Similarly, the effects of starvation, starvation in conjunction with

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sepsis (Fischer et al., 1996), proinflammatory cytokines (Pacitti et al. 1993), burn injury (Lohmann et al. 1998), advanced malignant disease (Inoue et al. 1995), and glucocorticoid excess can all increase glutamine transport and utilization within the liver (Bobe et al. 2002; Bode et al. 1995; Low et al. 1992). However, during metabolic acidosis glutamine flow is directed away from the liver and is significantly increased in the kidney (Welbourne and Oshi 1990). Under these conditions, hepatic ureagenesis is decreased, but renal ammoniagenesis is increased to facilitate H+ excretion.

5.3.

Regulation of Pyruvate Dehydrogenase Complex

The determining factor regulating formation of alanine and lactate from pyruvate is the activity of the pyruvate dehydrogenase complex. The pyruvate dehydrogenase (PDH) complex catalyzes the first irreversible reaction in the mitochondrial oxidation of glucose (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). Pyruvate undergoes oxidative decarboxylation by the PDH complex in the presence of NAD+ and CoA to form acetyl-CoA, NADH and CO2 (Figure 5.5). As such the PDH complex is the primary determinant of glucose oxidation in animal cells (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). The regulation of this complex is important to glucose homeostasis as it links glycolysis to tricarboxylic acid cycle and ultimately to ATP production by the mitochondria. Oxidation of pyruvate results in the depletion of body glucose carbon sources because glucose cannot be synthesized from acetyl-CoA and the PDH reaction is physiologically irreversible (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). Thus, the activity of the PDH complex determines whether pyruvate is oxidized to CO2 and H2 O or converted to lactate via lactate dehydrogenase or to alanine via alanine transaminase. Flux through the pyruvate dehydrogenase complex is tightly controlled through several mechanisms (Figure 5.6). The activity of the PDH complex is controlled both by end-product inhibition and by reversible phosphorylation (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). In vivo, the cycle of phosphorylation and dephosphorylation is accepted as the major form of regulation of the PDH complex, and the predominant mechanism controlling glucose oxidation in other conditions such as starvation or diabetes (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). The PDH complex is inactivated by phosphorylation, catalyzed by a PDH kinase (PDK), while a PDH phosphatase dephosphorylates the PDH complex and reactivates the enzyme complex (Harris et al. 2002; Holness and Sugden 2003; Randle et al. 1984; Randle 1998; Tsai et al. 1973). The PDH kinase and the PDH phosphatase are both active, and the relative activities of each of these two competing enzymes determines the proportion of PDH in the active, dephosphorylated form (PDHa ). Decreased flux through the

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Figure 5.5. Individual reactions catalyzed by subunits of pyruvate dehydrogenase complex.

PDH complex results from an increased proportion of inactive (phosphorylated) PDH complex (no change in total PDH activity) (Berger et al. 1976; Randle et al. 1984; Randle et al. 1988; Sugden et al. 1989). Inactivation of PDH complex limits oxidation of pyruvate derived from glycolysis and, hence, promotes increased lactate and alanine production. Accelerated oxidation of fatty acids raises the mitochondrial [acetylCoA]/[CoA] and [NADH]/[NAD+ ] concentration ratios, thereby stimulating the PDH kinase reaction (Hutson and Randle 1978; Kerbey et al. 1976; Vary et al. 1986). In other conditions, the PDHa may be increased in vitro by decreasing the fatty acid concentration in fed animals (Randle et al. 1988; Sugden et al. 1989), or in hearts by inhibiting fatty acid oxidation with 2-tetradecylglycidate (Caterson et al. 1982) or ranolazine (Clark et al. 1993; Clark et al. 1996). This mechanism represents a rapid onset/offset mechanism for short-term adjustments to the availability of fatty acids (Randle et al. 1988; Sugden et al. 1989). In starvation and diabetes, the short-term effects of oxidation of fatty acids to decrease the PDHa are supplemented by a more long-term mechanism involving

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Figure 5.6. Control of pyruvate dehydrogenase complex by phosphorylation/ dephosphorylation mechanism.

a stable increase in the activity of the PDH kinase (PDK). The activation of the PDK is observed under in vitro conditions where fatty acid oxidation does not occur. PDK activity is enhanced under these conditions by two mechanisms during starvation and diabetes. First, the specific activity of PDH kinase intrinsic to the complex is increased (Kerbey et al. 1979; Kerbey et al. 1984). Second, the abundance of the kinase is also increased (Jones and Yeaman 1991; Kerbey and Randle 1982). Indeed, PDK exists as genetically distinct isozymes (PDK1, PDK2, PDK3 and PDK4) (Bowker-Kinley et al., 1998; Gudi et al., 1995; Harris et al., 1995; Popov et al., 1993; Popov et al., 1994; Rowles et al., 1996). All members of this family contain conserved motifs that presumably form the kinase domain. The expression of these PDK isoenzymes occurs in a tissue-specific manner (BowkerKinley et al., 1998; Wu et al., 1988). The mRNA for PDK1 is localized almost exclusively in heart and pancreatic islets. PDK3 is limited to kidney, brain and testis. The mRNA for PDK2 was present in all tissues tested but its abundance was low in spleen and lungs. The mRNA for PDK4 was predominantly expressed in skeletal muscle and heart. The specific activities of the isoenzymes varies 25-fold from 50 nmol/min/mg for PDK2 to 1250 nmol/min/mg for PDK3

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(PDK1 = 650 nmol/min/mg; PDK4 = 400 nmol/min/mg). The isoenzymes also vary in the ability of dichloroacetate (DCA) to inhibit their activities (PDK2 > PDK4 > PDK1 > PDK3; from most sensitive to DCA to least sensitive to DCA), in the ability to be activated by NADH (PDK4 > PDK1 = PDK2 > PDK3), and in their ability to be activated by acetyl-CoA (PDK2 > PDK1 > PDK4 = PDK3). In vivo, overexpression of PDK2 inactivates PDC in cardiac muscle (Vary et al unpublished data). PDK4 is upregulated in most tissues during starvation, hormonal imbalances (diabetes, hyperthyroidism) and insulin resistance (Harris et al. 2002; Holness et al. 2002b; Holness and Sugden 2003; Randle 1998; Wu et al. 1999), but not aging (Moreau et al. 2004). The factors that appear to regulate PDK4 include fatty acid oxidation and adequate insulin signaling. The signaling pathways responsible for the changes in PDK4 expression remain unresolved. However, PDK4 expression is either directly or indirectly regulated by peroxisome proliferatorsactivated receptor alpha (PPAR-). PPAR- deficiencies limit the upregulation of PDK4 in liver and kidney during starvation. Administration of WY-14,643, a selective agonist for the peroxisome proliferator-activated receptor-alpha (PPAR-), induced large increases in pyruvate dehydrogenase kinase activity, PDK4 protein, and PDK4 mRNA in gastrocnemius muscle from wild–type but not in PPAR--null mice (Wu et al. 1999; Wu et al. 2001). Because long-chain fatty acids activate PPAR- endogenously, increased levels of these compounds in starvation and diabetes may signal increased expression of PDK4 in skeletal muscle. However, there is some controversy regarding the role of PPAR-alpha in regulating PDK4 expression in skeletal muscle and heart (Holness et al. 2002a; Holness et al. 2002b; Spriet et al. 2004). Whereas, activation of PPAR-alpha in gastrocnemius, a muscle composed primarily of fast, glycolytic fibers, increases PDK4 expression, the same effect is not observed in oxidative skeletal muscles. Likewise, skeletal muscle PDK-4 mRNA was progressively augmented over the course of a 40 h fast in humans, whereas PPAR-alpha protein and forkhead homolog in rhabdomyosarcoma (FKHR) mRNA abundance were unaffected by the fast (Spriet et al. 2004). The changes in PDK-4 expression and PDH activity did not coincide with increases in the transcriptional activators PPARalpha and FKHR. PDK4 protein expression in oxidative skeletal muscle and cardiac muscle appears regulated by a fatty acid-dependent mechanism that is not obligatorily dependent on signaling via PPAR-alpha, but may involve declines in plasma insulin concentrations. In this regard, PDK activity and the abundance of the PDK isoform 4 protein and mRNA become elevated prior to overt diabetes, but concomitant with rises in plasma fatty acids in animals that become diabetic spontaneously (Bajotto et al. 2004). Indeed, effects of insulin on PDK activity have shown that there are two phases to the insulin effect on PDC complex (Feldhoff et al. 1993). Acute activation of the PDH complex does not appear to result from lower PDK activity but rather correlates with an insulin-mediated lowering of plasma free fatty acids. However, with daily insulin therapy in diabetes, activation of PDH results from a decreased PDH kinase activity. In cardiac muscle, PPAR-alpha activation lowers glucose oxidation

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mainly by decreasing the flux of pyruvate through PDC due to negative feedback inhibition of PDC by fatty acid oxidation reaction products rather than by the phosphorylation of the PDC complex.

5.3.1. PDC in Regulation of Gluconeogenesis Maintenance of plasma glucose concentrations is achieved through glycogenolysis and by gluconeogenesis. Gluconeogenesis is the metabolic process by which glucose is formed from non-carbohydrate precursors. The process converts three carbon fragments derived primarily from lactate, glycerol and amino acids. The carbon source for gluconeogenesis arises from several sources including lactate, alanine, cysteine, and serine; substances released from peripheral organs in response to inhibition of PDC and accelerated proteolysis (Figure 5.3) (Vary 1999). The PDC lies at the crossroads of utilization of pyruvate for energy production and gluconeogenesis. Inactivation of PDC secondary to increased PDK activity promotes gluconeogenesis in starvation or diabetes by limiting oxidation of pyruvate thereby conserving three-carbon substrates (Vary 1999). This helps maintain glucose levels during starvation, but is detrimental in diabetes by inducing sustained hyperglycemia. The hallmark responsible for the transition of the liver and kidney to gluconeogenesis is lowering of plasma insulin concentrations or development of insulin resistance resulting in a sustained elevation in plasma lipids. Lipids act to lower PDC activity through increased acetyl-CoA/CoA ratios and expression of PDK4. The lipid-activated transcription factor, peroxisome proliferator-activated receptor-alpha (PPAR-), plays a pivotal role in the cellular metabolic response to fatty acids. In liver, decreased concentrations of insulin and increased concentrations of fatty acids and glucocorticoids promote PDK4 gene expression in starvation and diabetes through PPAR- (Huang et al. 2002). The decreased level of insulin is likely responsible for the increase in PDK2 mRNA levels in starvation and diabetes. Renal protein expression of PDK4 was only marginally induced by fasting in PPAR alpha null mice (Sugden et al. 2001). Specific up-regulation of PDK4 protein expression in starvation, by maintaining PDC activity relatively low, facilitates pyruvate carboxylation to oxaloacetate and therefore entry of acetyl-CoA derived from fatty acid betaoxidation into the TCA cycle, allowing adequate ATP production for brisk rates of gluconeogenesis. Indeed, activation of PDH by dichloroacetate (DCA) decreases peripheral release of alanine and lactate, thereby interrupting the Cori and alanine cycles and reducing the availability of three-carbon precursors for gluconeogenesis. DCA inhibits hepatic triglyceride and cholesterol biosynthesis. Administration of DCA in patients with non-insulin-dependent diabetes markedly reduces circulating very-low-density lipoprotein cholesterol and triglyceride concentrations (Stacpoole and Greene 1992). Unfortunately, DCA is metabolized to glyoxylate, which is converted to oxalate and, in the presence of adequate thiamine levels, to other metabolites which are toxic to humans (Stacpoole et al.

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1990). Hence, chronic use of DCA for diabetes mellitus or hyperlipoproteinemias is limited by its neurologic and other forms of toxicity.

5.3.2. PDH Phosphatase Activation of the PDH complex occurs through dephosphorylation catalyzed by two genetically different isozymes of pyruvate dehydrogenase phosphatase, PDP1c and PDP2c. PDP1c and PDP2c display marked biochemical and cellular distribution differences (Huang et al. 2003; Karpova et al. 2003). PDP1c is highly expressed in rat heart, brain, and testis and is detectable but less abundant in rat muscle, lung, kidney, liver, and spleen, whereas PDP2 is abundant in rat kidney, liver, heart, and brain and is detectable in spleen and lung (Huang et al. 2003). The catalytic subunit of pyruvate dehydrogenase phosphatase 1 (PDP1c) is a magnesium-dependent protein phosphatase heterodimer that regulates the activity of mammalian pyruvate dehydrogenase complex. The activity of PDP1c strongly depends upon the simultaneous presence of calcium ions and the E2 component of PDC. The stimulatory effect of E2 on PDP1c can be partially mimicked by a monomeric construct consisting of the inner lipoyl-bearing domain and the E1-binding domain of E2 component (Chen et al. 1996). This strongly suggests that the E2-mediated activation of PDP1c largely reflects the effects of co-localization and mutual orientation of PDP1c and E1 component facilitated by their binding to E2. Ca2+ appears to aid in the association of PDP1c with E2 bound, phosphorylated PDH (Chen et al. 1996). Site directed mutagenesis reveals that residues Asp54, and Asp347 contribute to the binuclear metal-binding center, and Asn49 contributes to the phosphate-binding sites (Karpova et al. 2004). The regulatory subunit of PDP (PDPr) lowers the sensitivity of the catalytic subunit of PDP1c to Mg2+ (Yan et al. 1996). In contrast, the activity of PDP2c displays little, if any, dependence upon either calcium ions or E2 (Karpova et al. 2003). Instead, PDP2c is sensitive to the biological polyamine spermine, which, in turn, has no effect on the enzymatic activity of PDP1 (Huang et al. 1998). Like PDP1, PDP2 is a Mg2+ -dependent enzyme, but its sensitivity to Mg2+ ions is almost 10-fold lower than that of PDP1. Spermine increases the sensitivity of PDP2c to Mg2+ , apparently by interacting with PDPr (Yan et al. 1996). Furthermore, PDP2c does not appreciably bind to PDC under the conditions when PDP1c exists predominantly in the PDC-bound state. Both PDP1c and PDP2c can efficiently dephosphorylate all three phosphorylation sites located on the alpha chain of the E1 component. For PDC phosphorylated at a single site, the relative rates of dephosphorylation of individual sites are: site 2 > site 3 > site 1. Phosphorylation of sites 2 or 3 in addition to site 1 does not have a significant impact on the rates of dephosphorylation of individual sites by PDP1c, suggesting a random mechanism of dephosphorylation. In contrast, a significant decrease in the overall rate of dephosphorylation of pyruvate dehydrogenase by PDP2c occurs under these conditions, suggesting that the mechanism of dephosphorylation of PDC phosphorylated at multiple sites by PDP2c is not purely random. Starvation and

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diabetes lower PDP2 mRNA abundance, PDP2 protein content, and PDP activity in rat heart and kidney. Refeeding and insulin treatment effectively reversed these effects of starvation and diabetes, respectively on PCP (Huang et al. 2003). These marked differences in the site-specificity displayed by PDP1c and PDP2c may be important under conditions (e.g. starvation and diabetes) associated with increases in phosphorylation of sites 2 and 3 of pyruvate dehydrogenase and provide a plausible mechanism for the observed changes in phosphorylation of different sites of the PDH complex upon reactivation (Randle et al. 1984).

5.3.3. Role of Pyruvate Dehydrogenase Complex in Sepsis-Induced Hypermetabolism Severe trauma and sepsis initiate a pattern of physiologic and metabolic adaptations characterized by an increase in oxygen consumption and alterations in carbohydrate, fat, and protein metabolism. Virtually every organ system of the body becomes affected displaying recognizable metabolic adjustments. The host’s metabolic response to sepsis is associated with the rapid breakdown of the body’s reserves of protein and carbohydrates, and an increased dependence upon fatty acids. Sepsis shifts interorgan substrate fluxes causing an exaggerated response in the normal physiologic mechanisms responsible for maintenance of whole-body metabolic homeostasis. The degree of metabolic dysfunction is ultimately indicative of the extent of organ dysfunction induced by the invading organism or traumatic injury. Two of the earliest and most recognizable manifestations of organ dysfunction in sepsis are a massive excretion of urea resulting in a large negative nitrogen balance and alterations in glucose kinetics resulting in increased rates of glucose appearance (150–200%) and metabolic clearance. As such, sepsis represents the most common causes of lactic acidosis in the presence (59%) or absence (49%) of shock (Stacpoole et al. 1994). Sepsis increases the rate of glucose appearance by stimulating hepatic glucose production (Keller et al. 1982; Lang et al. 1987; Vary et al. 1988b). Of the gluconeogenic precursors, lactate accounts for 60–70% of the glucose carbon used for gluconeogenesis. Net lactate extraction by the liver is increased 2–3 fold following severe infection (Dahn et al. 1987; McGuinness et al. 1995; Wilmore et al. 1980). An increased rate of glucose production indicates that hepatic lactate utilization is accelerated in sepsis. Thus, a constant supply of gluconeogenic precursors must be maintained for the enhanced and persistent rates of gluconeogenesis during sepsis. The release of lactate and alanine from skeletal muscle is increased, indicating both the Cori cycle and Felig cycle are providing adequate precursors for gluconeogenesis in sepsis. Skeletal muscle also represents a major source for the production of gluconeogenic substrates. In contrast to protein catabolism, altering substrate delivery to the liver by modulating skeletal muscle glucose metabolism by treatment with dichloroacetate to activate PDC short-circuited the characteristic derangements in glucose kinetics during sepsis. Thus, the accelerated glucose appearance and glucose clearance appears dependent upon increased

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lactate and alanine production by skeletal muscle during sepsis. Elevated plasma lactate concentrations are prognostic indicators of illness severity and/or death in septic patients (Cerra 1989; Clark et al. 1996; Siegel et al. 1990; Vary et al. 1988a). Skeletal muscle, by virtue of its mass, represents the major source of gluconeogenic substrates. Sepsis dramatically enhances the peripheral utilization of glucose, such that the glucose metabolic clearance rate is increased over 2-fold compared with non-septic animals (Lang et al. 1990; Vary et al. 1988b). Uptake of glucose by skeletal muscle is enhanced by systemic infection. Stimulation of glucose uptake is not associated with increased deposition of glycogen (Vary et al. 1995) but instead there is an acceleration of glycolysis with the subsequent release lactate and alanine from skeletal muscle (Gump et al. 1975; Shaw et al. 1985; Vary et al. 1988b; Vary and Murphy 1989). The increased lactate production by skeletal muscle provides the necessary gluconeogenic precursors for maintenance of sustained rates of gluconeogenesis in sepsis. Furthermore, lactate production most likely exceeds its utilization, thereby accounting for elevated plasma lactate concentrations in sepsis. A strong correlation is obtained between arterial lactate concentration and the rate of glucose appearance or rate of recycling of glucose carbon (Lang et al. 1987) implying that circulating lactate concentrations are an important determinant of gluconeogenesis. Lactate arises from the reduction of pyruvate. Lactate production occurs whenever the rate of pyruvate production from glycolysis or other sources [malic enzyme or pyruvate carboxylase] exceeds glucose oxidation by the mitochondria. Therefore, glucose oxidation should decrease resulting in increased gluconeogenesis, with a higher rate of diversion of pyruvate to lactate being important physiologically in the regulation of plasma lactate concentrations. In septic patients, whole-body glucose oxidation is inhibited and glucose turnover and recycling is increased (Jahoor et al. 1989; Shaw et al. 1985). Accelerated lactate release by the forearm of post-surgical patients occurs at a time when glucose oxidation is depressed, indicating such a scenario occurs in vivo during sepsis (Brandi et al. 1993). Mitochondria isolated from skeletal muscle of septic rats have a defect in their ability to oxidize pyruvate, but not other substrates, including fatty acids (MelaRiker et al., 1992). This implies that there is specific sepsis-induced inhibition of the PDC. The inhibition of PDH complexes is a consequence of an increased acetyl-CoA/CoA ratio, secondary to augmented fatty acid oxidation (Vary et al. 1986). The importance of inhibiting the PDH complex in controlling plasma lactate concentrations is indicated by two observations. First, a negative linear correlation exists between plasma lactate concentrations and skeletal muscle PDHC in the active dephosphorylated form (Vary 1996). Second, activation of the PDH complex with administration of dichloroacetate (DCA), an noncompetitive inhibitor of the PDH kinase, normalizes muscle and plasma lactate and alanine concentrations and inhibits skeletal muscle lactate and alanine production during sepsis (Lang et al. 1987; Vary et al. 1988b; Vary et al. 1989a; Vary et al. 1989b; Vary 1996).

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Does the ability of DCA to activate the PDH complex, thereby enhancing pyruvate oxidation and subsequently lowering skeletal muscle lactate and alanine concentrations and production, modulate glucose kinetics during sepsis? After injection of DCA into septic rats, the elevated rates of glucose appearance (Lang et al. 1987) and turnover were reduced (Vary et al. 1988b). Furthermore, the sepsis-increased glucose metabolic clearance rate is normalized (Vary et al. 1988b). Hence, activation of the PDH complex successfully reversed or attenuated the sepsis-induced increases in glucose rate of appearance, glucose carbon recycling, and glucose metabolic rate. These observations indicate enhanced rates of gluconeogenesis are dependent in part upon the sustained delivery of precursors from skeletal muscle from muscle to liver. Thus, stimulated glucose uptake and glycolysis coupled with an inhibition of PDH complex in skeletal muscle and subsequent release of lactate and alanine drives the accelerated rates of gluconeogenesis during sepsis.

5.3.4. Potential Beneficial Therapeutic Effects of Pyruvate The inhibition of the PDC and the subsequent hyperlactatemia and hyperalaninemia under a variety of conditions has prompted the notion of using inhibitors of the PDHK to modify metabolic derangements with the assumption that this would lead to an increased beneficial effect. Indeed, dichloroacetate has been touted as a useful therapeutic modality in treating hyperlactatemia (Vary et al. 1988a) and in animal models of malaria prolongs survival (Holloway et al. 1995). Activation of PDC improves heart function (Kline et al. 1997) and survival following hemorrhagic shock (Granot and Steiner 1985). Unfortunately, a multicenter double-bind randomized trial failed to show improvement in hemodynamics or survival following activation of PDC despite lowering plasma lactate concentrations and increasing plasma arterial pH (Stacpoole et al. 1992). One potential reason for such a result was that both hemodynamic and non-hemodynamic (metabolic) underlying causes were present in the patient population many of which independently predicted survival and most of who were refractory to standard care. Alternatively, elevating the cellular content of pyruvate also inhibits the PDHK. In addition, pyruvate may act as an endogenous antioxidant and free radical scavenger. Recognition that pyruvate acts both as an effective activator of PDC and ROS scavenger has led investigators to use this compound as a potential therapeutic agent for the treatment of various pathologic conditions (Liedtke et al. 1976; Mochizuki and Neely 1980; Mongan et al. 1999; Slovin et al. 2001). The usefulness of pyruvate as a therapeutic agent is abrogated by its very poor stability in solution. Aqueous solutions of pyruvate rapidly undergo an aldol-like condensation reaction to form 2-hydroxy-2-methyl-4-ketoglutarate (parapyruvate), a compound that is a potent inhibitor of a critical step in the mitochondrial tricarboxylic acid cycle (Montgomery and Webb 1956). Ethyl pyruvate, a stable derivative of pyruvic acid in solution, circumvents the problems relating to stability of pyruvate in solution (Sims et al. 2001).

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Original reports in the literature indicated that ethyl pyruvate could attenuate deleterious effects of chemically-induced oxygen free radical damage in the lens (Varma et al. 1998). Subsequently, treatment with ethyl pyruvate under a variety of conditions ameliorates much of the structural and functional damage caused by the insult. Treatment with ethyl pyruvate solution could improve survival in rodent models of hemorrhagic shock or myocardial ischemia (Tawadrous et al. 2002; Woo et al. 2004), acute endotoxemia and bacterial peritonitis (Fink 2003a; Ulloa et al. 2002), and sepsis-induced renal failure (Miyaji et al. 2003). The protective actions may be related to metabolic (increased ATP production from pyruvate) and non-metabolic (anti-inflammatory) mechanisms (Fink 2003b) that down-regulate a number of proinflammatory genes. The anti-inflammatory effects of ethyl pyruvate may be mediated through inhibition of NF-kappaB signaling to p65 by binding ethyl pyruvate to p65 at Cys(38) (Han et al. 2005).

5.3.5. Role of Pyruvate Dehydrogenase Complex in Prostate Cancer Prostate cancer is the most common cancer detected in American men and the second leading cause of cancer-related deaths in the United States. Most prostatic cancers are detected in asymptomatic men who are found to have focal nodules or areas of induration within the prostate at the time of digital rectal exam or more recently through screening for the prostate-specific antigen in the blood or transrectal ultrasound. The PSA that alone is not specific for cancer but is more commonly associated with benign prostate hyperplagia which occurs frequently in the same age groups as prostate cancer patients. The prostate gland secretes a thin, milky fluid that contains calcium, citrate, phosphate, a clotting enzyme, and a profibrinolysin and eventually becomes incorporated, with secretions from other tissues, into semen. The prostate gland has the unique function of accumulating and secreting extraordinarily high levels of citrate. The prostate secretory epithelial cells synthesize citrate which, due to a limiting mitochondrial (m-) aconitase, accumulates rather than being oxidized. Thus citrate becomes essentially an end product of metabolism in prostate. For continued net citrate production, a continual source of oxaloacetate (OAA) and acetyl CoA is required. Glucose via pyruvate oxidation provides the source of acetyl-CoA. Citrate production is regulated by testosterone and/or by prolactin (Costello et al. 2000). Both hormones selectively regulate the level and activity of pyruvate dehydrogenase E1 alpha (E1a), thereby regulating the availability of acetyl CoA for citrate synthesis. This process is under regulatory control by testosterone and prolactin, which serve to modulate the expression of the genes associated with these mitochondrial enzymes. The regulation of these genes by testosterone and prolactin is specific for prostate cells, and appears to be mediated through signaling pathways involving protein kinase C. Of interest to the pathogenesis of prostate cancer, the ability of the prostate to secrete citrate is retarded in cancer cells whereas increased citrate production is characteristic of benign prostate

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hyperplagia. These disease entities therefore represent a unique regulation of mitochondrial function in that there is regulation of the gene expression of key regulatory enzymes that are under hormonal control.

5.4.

Branched-Chain Amino Acid Dehydrogenase

Branched chain amino acids (BCAA) (leucine, valine, isoleucine) are essential amino acids whose major function in the body may not reside with energy production but rather as a nutrient signal regulating protein metabolism. Of the branched-chain amino acids, leucine is well suited as a nutrient signal for several reasons. First, leucine is the most potent of the branched chained amino acids with regard to accelerating protein synthesis. Second, it cannot be synthesized de novo by mammals. Thus, changes in the plasma leucine concentrations must arise from nutrition, tissue breakdown or changes in the rate of oxidation or loss. Third, it is the most abundant essential amino acid in dietary protein and thus is a crucial nutrient following ingestion of a protein meal. Fourth, once taken up by the gut, leucine cannot be directly metabolized by the liver, which lacks the branched-chain amino transferase (BCATm), the mitochondrial enzyme responsible for removal of the nitrogen. This is in contrast to the liver’s capacity to initiate the catabolism of other amino acids. In contrast to liver, most other peripheral tissues have excess capacity (BCATm) to catalyze this first step. This may facilitate postprandial rises in plasma leucine concentrations as leucine would not be cleared in first pass by the liver. BCAA catabolism entails two enzymatic steps common to all BCAAs followed by subsequent steps that are dependent upon BCAA being utilized. The first step involves reversible transamination of the BCAA with -ketoglutarate to form branched chain alpha-keto acids (BCKAs) and glutamate (Figure 5.1). The glutamate formed is then transaminated with pyruvate to form alanine and regenerate -ketoglutarate or the glutamate is converted to glutamine. Following transamination, BCKAs undergo irreversible oxidative decarboxylation. The BCAA catabolic enzymes are distributed widely in body tissues and, with the exception of the nervous system, all reactions occur in the mitochondria of the cell. Transamination provides a mechanism for dispersing BCAA nitrogen according to the tissue’s requirements for glutamate and other dispensable amino acids. The intracellular compartmentalization of the branched-chain aminotransferase isozymes (mitochondrial branched-chain aminotransferase, cytosolic branched-chain aminotransferase) impacts on intra- and interorgan exchange of BCAA metabolites, nitrogen cycling, and net nitrogen transfer. The branched-chain ketoacid dehydrogenase (BCKDC) multienzyme complex catalyzes the rate-limiting step and is the first irreversible step in leucine oxidation. As such, the BCKDC is the most important regulatory enzyme in the catabolic pathways of the BCAAs. The reaction converts branched-chain keto acids [-ketoisocaproate acid (KIC)] to CoA derivatives (such as isovaleryl CoA). The BCKDC is a multi-enzyme complex. It contains a branched-chain keto

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acid decarboxylase (E1, which has an 22 structure with a covalently bound thiamine pyrophosphate cofactor), dihydrolipoyl transacylase (E2, which has a covalently bound lipoic acid), and the flavin-linked dihydrolipoyl dehydrogenase (E3) (for review see Ref. (Harper et al. 1985)). Activity of the complex is controlled by covalent modification with phosphorylation of its branched-chain alpha-ketoacid dehydrogenase subunits by a specific kinase [branched-chain kinase (BDK)] causing inactivation and dephosphorylation by a specific phosphatase [branched-chain phosphatase (BDP)] causing activation. Tight control of BCKDC activity is important for conserving as well as disposing of BCAAs. Phosphorylation of the complex occurs when there is a need to conserve BCAAs for protein synthesis; dephosphorylation occurs when BCAAs are present in excess. The relative activities of BDK and BDP set the activity state of BCKDC. BDK activity is regulated by alpha-ketoisocaproate inhibition and altered level of expression. The -keto acid of leucine, KIC, is thought to inhibit the BCKD kinase, promoting activation of BCKD complex. BCKD kinase phosphorylates the E1 subunit of BCKD at two sites, termed sites 1 and 2 (Harris et al. 1986; Paxton et al. 1986). Phosphorylation at site 1 (Ser293 ) inhibits BCKD activity, whereas phosphorylation at site 2 is silent (Zhao et al. 1994). The BCKDH kinase is responsible for inactivation of the complex by phosphorylation, and the activity of the kinase is inversely correlated with the activity state of the BCKDH complex, which suggests that the kinase is the primary regulator of the complex. Ligands for peroxisome proliferator-activated receptor-alpha (PPAR-) in rats caused activation of the hepatic BCKDH complex in association with a decrease in the kinase activity, which suggests that higher rates of fatty acid oxidation upregulates the BCAA catabolism (Shimomura et al. 2004b). Less is known about BDP but a novel mitochondrial phosphatase was identified recently that may contribute to the regulation of BCKDC. Whereas BCAA transamination occurs primarily in extrahepatic tissues, liver is thought to be the major site of BCAA oxidation. It has been shown that kinase activity and expression are affected by nutritional states imposed by lowprotein diet feeding, starvation, and diabetes. Evidence has also been presented that certain hormones, particularly insulin, glucocorticoid, thyroid hormone and female sex hormones, affect the activity and expression of the kinase. The findings indicate that nutritional and hormonal control of the activity and expression of branched-chain alpha-keto acid dehydrogenase kinase provides an important means of control of the activity of the branched-chain alpha-keto acid dehydrogenase complex, with inactivation serving to conserve branched-chain amino acids for protein synthesis in some situations and activation serving to provide carbon for gluconeogenesis in others (Shimomura et al. 2001). In rats fed a standard laboratory diet (∼20% protein), which is more than sufficient to meet the animal’s protein requirements, the liver enzyme is essentially active (Block et al. 1990; Gillim et al. 1983; Miller et al. 1988). Only low-protein diets had a significant impact on activity state. Dietary protein restriction results in increased expression of BCKD kinase (Popov et al. 1995), and high levels of kinase were found with BCKD, leading to increased steady-state levels of phosphorylated

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E1 and conservation of essential BCAA. Once dietary protein exceeded the requirement, little kinase was found with BCKD in the PEG pellet. Decreasing dietary protein lowers BCKD kinase mRNA and activity (Popov et al. 1995). Lower concentrations of E2 and E1 were also observed in the low-protein diet in addition to the changes in BCKD kinase (Lynch et al. 2003a and b). Hence, dietary intake of leucine can regulate its metabolism in the mitochondria.

5.4.1. Maple Syrup Urine Disease (Branched Chain Ketoaciduria) Maple syrup urine disease (MSUD) is a disorder caused by a deficiency of the branched-chain alpha-ketoacid dehydrogenase complex (BCKD). The defect leads to an accumulation of both the branched chain amino acids and branched chain -keto acids. Indeed, the molar ratio of leucine to alanine is elevated 10- to 20-fold in this disease. The disease gets its name from the maple syrup odor of the urine which is caused by the excretion of excessive amounts of -keto3-methylvalarate in the urine of affected patients, producing an odor similar to that of maple syrup. MSUD is characterized by psychomotor retardation, feeding problems, and a maple syrup odor of the urine initially diagnosed in newborns. Mutations in the genes encoding BCKD complex components E1, E1, E2 and E3 mapped to human chromosomes 19q13.1-q13.2, 6p22-p21, 1p31, and 7131q32, respectively are the causative agents. The mode of inheritance is autosomal recessive with an incidence of MSUD of approximately 1 in 180,000 births in the United States. The incidence increases to as high as 1 in 200 births in populations that have significant inbreeding communities, such as the Mennonites living in Pennsylvania. There are five distinct phenotypes of MSUD: classical, intermittent, intermediate, thiamine-responsive, and E3-deficient. Classical MSUD typically presents in newborns and represents the most common form of the disease. Newborns typically develop ketonuria within 48 hours of birth and present with irritability, poor feeding, vomiting, lethary and dystonia. By four days of age, neurologic abnormalities include alternating lethary and irritability, dystonia, apnea, seizures, and indications of cerebral edema. The initial symptoms may not develop until the infant is four to seven days of age, depending upon the feeding regimen and breastfeeding may delay onset of symptoms to the second week. Dietary therapy to promote normal growth and development and prompt treatment of episodes of acute metabolic decompensation constitute the cornerstone of management of these patients. Despite careful dietary management, episodes of metabolic intoxication may occur in affected older infants or children by increased net endogenous protein catabolism induced by intercurrent illness, exercise, injury, surgery, cancer or fasting. Clinical manifestations include epigastric pain, vomiting, anorexia and muscle fatigue. Neurologic signs may include hyperactivity, sleep disturbance, stupor, decreased cognitive function, dystonia and ataxia. The best outcomes occur in newborns, where therapy is begun before they become symptomatic or are treated rapidly after symptoms

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develop. Acute metabolic decompensation can result in brain injury and requires prompt treatment to avoid neurologic sequelae. Even with aggressive treatment MSUD can be fatal in the newborn period or during decompensation. The intermediate, intermittent and thiamine-responsive forms of MSUD may present at any time during infancy or childhood and appear to be brought on by episodes of stress. Intermediate MSUD is a rare disorder caused by mutations in the E1 component of BCKD but in contrast to the classical MSUD, residual BCKD activity typically is 3 to 30% of normal. Patients may become symptomatic at any age, when the flux through BCATm exceeds flux through BCKD. The intermittent form of the disease is the second most common type of MSUD. Affected patients have normal growth and development. They typically present with ketoacidosis during episodes of catabolic stress in childhood, including intercurrent illnesses such as otitis media. Signs of neurotoxicity develop, including ataxia, lethargy, seizures and coma. Death may occur without appropriate recognition and treatment. Thiamine-responsive MSUD is a rare phenotype that has been associated with mutations in the E2 component, the subunit that requires thiamine as a cofactor for complete enzymatic activity (Figure 5.7). Clinical presentation is similar to the intermediate form. In general, symptoms in affected patients are not completely resolved with thiamine supplementation alone, necessitating dietary restriction of branched-chain amino acids to achieve metabolic control. Mutations in the gene encoding the E3 component of BCKD (Figure 5.7) result in E3-deficient MSUD. Because the E3 component of BCKD is also present in the PDH complex (Figure 5.5) and -ketoglutarate dehydrogenase, affected patients show symptoms of combined deficiencies of BCKD and the pyruvate complexes including lactic acidosis superimposed on MSUD.

5.4.2. Role of Mitochondria in Cell Signaling Pathways Activated by Leucine Leucine is a direct acting nutrient signal that regulates protein synthesis in a number of tissues. The effects on protein synthesis are brought about, at least in part, by activation of cell-signaling pathways. These pathways are regulated by adapter proteins, such as RAPTOR (Hara et al. 2002; Kim et al. 2002), and by phosphorylation/dephosphorylation cascades (e.g., mTOR to ribosomal protein S6 kinase-1 (S6K1) to S6 and mTOR to 4E-BP1 to eIF4E·eIF4G) that, in turn, modulate factors involved in initiation of protein synthesis (Figure 5.8). In striated muscle, the mTOR signaling pathway appears to play a role in stimulation of protein synthesis by growth promoting anabolic hormones. Two well known substrates of mTOR, namely 70 kDa ribosomal protein S6 kinase (S6K1) and the translational repressor, eukaryotic initiation factor 4E binding protein 1 (4E-BP1), have been implicated in the regulation of protein synthesis. At least two signaling pathways appear to be important in leucine’s acute regulation of protein synthesis, a rapamycin-sensitive pathway involving mTOR and an unknown rapamycin-insensitive pathway that may or may not involve

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Figure 5.7. Individual reactions catalyzed by subunits of branched chain amino acid dehydrogenase complex.

mTOR. It is presently unclear exactly how leucine activates mTOR aside from the fact that the mechanism differs from that used by insulin. For example, no leucine receptor has been yet identified. In fact, it has not been unequivocally established whether effects of leucine are mediated by leucine or a metabolite derived

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Figure 5.8. Dual role of leucine in contributing to energy metabolism and control of signaling pathways involved in gene expression. mTOR – mammalian target of rapamycin; Glu – glutamate; -KG – -ketoglutarate; PKC – protein kinase C epsilon isoform; 4E-BP1 – eukaryotic initiation factor 4E binding protein 1; S6K1 – ribosomal protein S6 protein kinase 1; BCKD – branched chain amino acid dehydrogenase.

from leucine transamination such as -ketoisocaproate (-KIC) or leucine oxidation. Leucine’s effects on protein metabolism are self-limiting because leucine promotes its own disposal by an oxidative pathway, thereby terminating its positive effects on body protein accretion. A strong case can therefore be made that the proper leucine concentration in the various compartments of the body is critically important for maintaining body protein levels beyond simply the need of this essential amino acid for protein synthesis. In this regard, mitochondrial metabolism may be required for leucine activation of mTOR. Specifically, a metabolically-linked signal arising from activation of leucine metabolism in the mitochondria results in mTOR activation (McDaniel et al. 2002). A potential target cited was the branched chain ketoacid dehydrogenase (BCKD) complex. There are several compelling arguments in favor of the notion that BCKD kinase activities might provide a link between leucine signal and the activation of mTOR. The first is that -ketoisocaproate acid is a physiological inhibitor of BCKD kinase. The second is that the order of potency of Leu > Ile > Val

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and importance of leucine in the activation of mTOR signaling is similar to that of their respective keto acids’ ability to activate the BCKD complex via inhibition of the BCKD kinase. A third observation that supports this potential mechanism is that mTOR appears to be associated with mitochondria membrane in situ. One way to delineate this effect is to simultaneously examine the dose dependent effects of orally administered leucine on acute activation of S6K1 (an mTOR substrate) and phosphorylation of BCKD (Lynch et al. 2003a). Increasing doses of leucine given orally via gavage directly correlated with elevations in plasma leucine concentration. Phosphorylation of S6K1 (Thr389, the phosphorylation site leading to activation) in adipose tissue was maximal at a dose of leucine that increased plasma leucine approximately threefold. Changes in BCKD phosphorylation state required higher plasma leucine concentrations. Hence there exists a disconnection between the concentration of leucine required for activation of S6K1 system and phosphorylation of BCKD. Only at concentrations significantly higher than those required for activation of S6K1 is the phosphorylation state of BCKD lowered, allowing for oxidation of leucine to occur. Hence as the leucine signal becomes too high, there is a stimulation of leucine oxidation in peripheral tissue thereby lowering the plasma leucine concentration. Thus, leucine metabolism may only be important in regulating the magnitude and duration of the signal for mTOR activation. Leucine dependent stimulation of mRNA translation in vivo occurs in part, through a rapamycin-sensitive pathway (Anthony et al. 2000; Anthony et al. 2001). Rapamycin treatment, an inhibitor of the protein kinase mTOR, reduced the mTOR-dependent 4E-BP1 and S6K1 phosphorylation. However, leucine retained its ability to stimulate protein synthesis. Thus, inhibition of mTORmediated signaling was not capable of completely eliminating the leucinedependent stimulation of protein synthesis. An additional study examining oral leucine administration in diabetic rats demonstrated enhanced levels of protein synthesis in skeletal muscle without corresponding changes in 4E-BP1 or S6K1 (Anthony et al. 2002). Similarly rates of skeletal muscle protein synthesis in perfused hindlimbs can be enhanced by elevated leucine concentrations in the absence of phosphorylation changes of 4E-BP1 and S6K1. Collectively, this evidence underscores the ability of leucine to modulate translation initiation and protein synthesis through an mTOR-independent mechanism(s).

5.4.3. Branched Chain Amino Acid Metabolism in Liver Disease Hepatic encephalopathy is a neuropsychiatric syndrome occurring in patients with acute or chronic liver disease including cirrhosis. During the last 20 years there has been much interest in nutritional treatment for patients with advanced cirrhosis. Most studies have measured the potential benefit of nutritional supplements of dietary proteins, generic protein hydrolysates, or specific branchedchain amino acid (BCAA)-enriched formulas in regard to nutritional parameters

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and hepatic encephalopathy. Nutritional factors play a major role both in the pathogenesis as well as management of hepatic encephalopathy (HE). Physicians treating patients with chronic liver disease often restrict the intake of dietary protein to prevent a rise in blood ammonia levels. The role of protein restriction in patients with chronic hepatic encephalopathy (CHE) has been questioned recently as the efficacy of protein withdrawal in patients with CHE has never been subjected to a controlled trial. Evidence suggests that protein intake plays only a limited role in precipitating encephalopathy. In fact, measures taken to suppress endogenous protein breakdown are more effective than dietary restrictions in reducing the load of amino acids on the decompensated liver. A protein intake of less than 40 g per day, as has been indicated, contributes to a negative nitrogen balance, which along with increased endogenous protein catabolism, worsens encephalopathy. A positive nitrogen balance may have positive effects on encephalopathy. Rather, depressed plasma branched-chain amino acid (BCAA) levels, implicated in the pathogenesis of HE, also supervene in cirrhosis only when malnutrition is present as well. Therefore, the emphasis in the nutritional management of patients with CHE should not be on the reduction of protein intake (Srivastava et al. 2003). Eleven randomised trials (556 patients) assessing BCAA versus carbohydrates, neomycin/lactulose, or isonitrogenous control showed BCAA significantly increased the number of patients improving from hepatic encephalopathy at the end of treatment although there were methodological concerns (Als-Nielsen et al. 2003; Fabbri et al. 1996). Chemically-induced liver cirrhosis in rats raised the activity of hepatic BCKDH complex and decreased plasma BCAA and branched-chain -keto acid concentrations, suggesting that the BCAA requirement is increased in liver cirrhosis. Because the effects of liver cirrhosis on the BCKDH complex in human liver are different from those in rat liver, further studies are needed to clarify the differences between rats and humans. In the valine catabolic pathway, crotonase and beta-hydroxyisobutyryl-CoA hydrolase are very important to regulate the toxic concentration of mitochondrial methacrylyl-CoA, which occurs in the middle part of valine pathway and highly reacts with free thiol compounds. Both enzyme activities in human and rat livers are very high compared to that of the BCKDH complex. It has been found that both enzyme activities in human livers were significantly reduced by liver cirrhosis and hepatocellular carcinoma, suggesting a decrease in the capability to dispose methacrylyl-CoA (Shimomura et al. 2004a). The findings described here suggest that alterations in hepatic enzyme activities in the BCAA catabolism are associated with liver failure. Metabolic acidosis increases protein degradation resulting in muscle wasting and a negative nitrogen balance. The branched-chain amino acids serve as useful markers of these changes and their catabolism is increased in acidosis, particularly for the spontaneous acidosis associated with renal failure (Bailey 1998; Wang et al. 1997). As a result, the neutral nitrogen balance is compromised and malnutrition results. Glucocorticoids mediate these changes through the recently discovered ATP-dependent ubiquitin-proteasome pathway. Metabolic acidosis and glucocorticoids act in concert to stimulate branched-chain amino

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acid (BCAA) oxidation in adrenalectomized rats. In muscles of normal rats, metabolic acidosis increases the maximal activity of the rate-limiting enzyme, branched-chain alpha-ketoacid dehydrogenase (BCKAD) and a genetic response to catabolic conditions like uremia is implicated by concurrently higher levels of BCKAD subunit mRNA. Independent transactivation response elements to acidification or glucocorticoids were localized in the E2 promoter. In summary, catabolic responses to low extracellular pH and glucocorticoids include enhanced expression of genes encoding BCKAD subunits. (Price and Wang 1999).

5.5.

Summary

The regulation of the PDH and BCDH was originally established nearly 30 years ago as a means to help explain the control of carbohydrate and amino acid metabolism during transitions from feeding to starvation and during conditions involved in altered carbohydrate metabolism. These types of studies also delineated the role of glutamine in amino nitrogen metabolism in the body. In recent years, there has been renewed interest in both the structure, function and interaction of the components of the PDH and BCDH to define the molecular interactions responsible for control mechanisms. More importantly, there has been a growing body of evidence suggesting that these amino acids can function as key effector molecules altering the signaling cascades responsible for regulatory several intracellular systems responsible for synthesis and expression of specific gene products. Future studies will have to integrate the mechanisms of metabolism of pyruvate, glutamine and leucine with their roles in the controlling gene expression. Acknowledgements. This work was supported NIGMS Grant GM-39277 (TCV), NIAAA Grant AA12814 (TCV), NIDDK Grant DK053843 (CJL) and NIDDK Grant DK062880 (CJL) awarded by the National Institutes of Health.

References Abcouwer SF, Bode BP, Souba WW (1995) Glucocorticoids regulate rat glutamine synthetase expression in a tissue specific manner. J Surg Res 59: 59–65 Abcouwer SF, Norman J, Fink G, Carter G, Lustig RJ, Souba WW (1996) Tissue-specific regulation of glutamine symthetase gene expression in acute pancreatitis is confirmed by using interleukin-1 receptor knockout mice. Surgery 120: 255–263 Abcouwer SF, Lohmann R, Bode BP, Lustig RJ, Souba WW (1997) Induction of glutamine synthetase expression after major burn injury is tissue specific and temporally variable. J Trauma 42: 421–427 Als-Nielsen B, Koretz RL, Kjaergard LL, Gluud C (2003) Branched-chain amino acids for hepatic encephalopathy. Cochrane Database Systematic Rev 2: CD001939 Anthony JC, Yoshizawa F, Anthony TG, Vary TC, Jefferson LS, Kimball SR (2000) Leucine stimulates translation initiation in skeletal muscle of postabsorptive rats via a rapamycin-sensitive pathway. J Nutr 130: 2413–2419

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6 Amino Acids and the Mitochondria Nicola King

6.1.

Introduction and Summary

This chapter describes some of the important physiological functions of amino acids in the mitochondria and the alterations caused by specific pathologies. To some extent all of the featured items are dependent upon the movement of amino acids across the highly selective permeability barrier that is the inner mitochondrial membrane. The performance of this transport by specific carriers is the subject of the first section. Once inside the mitochondrial matrix the amino acids become involved in a bewildering number of critical metabolic pathways. The second section elaborates on two of the most significant namely: the malateaspartate shuttle essential for the transfer of reducing equivalents between the cytoplasm and the mitochondria; and the urea cycle, which is responsible for maintaining sub-toxic levels of ammonia. The final section covers the changes to mitochondrial amino acid metabolism that occur under different pathological conditions. In this case three examples have been chosen comprising ischemiareperfusion in heart, myocardial hypertrophy and the special relationship that exists between glutamine and cancer cells.

6.2. Amino Acid Transport Across the Mitochondrial Inner Membrane 6.2.1. Introduction When non-respiring mitochondria are suspended in isotonic solutions containing different amino acids they swell suggesting that these compounds can cross the mitochondrial inner membrane (Palmieri et al. 2001; Porter 2000). Amino acid flux through the mitochondrial inner membrane has also been measured directly using radiotracers via inhibitor stop techniques, oil- or Millipore- filtration (reviewed in LaNoue and Schoolwerth 1979; Passarella et al. 2003). These investigations have greatly magnified our knowledge of the transport characteristics of the various carriers (Figure 6.1), although surprisingly only a relatively small number of the genes and proteins responsible have been identified. 151

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Figure 6.1. Amino acid carriers in the inner mitochondrial membrane. Question marks indicate carriers that have been functionally characterized where the genes and proteins responsible are unknown. For the sake of clarity only the major substrates are shown. In contrast with those carriers, which have been identified at the molecular level, the names of the proteins and all the natural amino acid substrates are given.

6.2.2. Acidic Amino Acid Carriers The main acidic amino acid carriers of the mitochondrial inner membrane are the aspartate-glutamate exchangers and the glutamate carriers. Some of the most important characteristics of the aspartate-glutamate exchangers will be discussed later (section 6.3.2), because of their key role in the malate-aspartate shuttle. However, it is worth mentioning here that these carriers have narrow substrate specificities, i.e. the only other natural amino acid to be accepted is cysteinesulfinate (LaNoue and Schoolwerth 1979). This compound, which is important for sulfur homeostasis (Townsend et al. 2004), can be involved in a straight exchange for aspartate or in an electrogenic exchange for glutamate and a proton (LaNoue and Schoolwerth 1979). There are two aspartate-glutamate exchangers that have so far been identified, which are encoded by the genes, SLC25A12 and SLC25A13 (Palmieri et al. 2001). The encoded proteins named aralar 1 and aralar 2 (also called citrin) respectively display the typical features of the

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mitochondrial carrier family including EF hand Ca2+ binding motifs in their N terminals (4 in each protein) (Palmieri et al. 2001). Aralar 1 is most prevalent in heart, skeletal muscle and brain, whereas aralar 2 is expressed in many tissues but highest levels are in liver (Palmieri et al. 2001). Mutations in the gene encoding aralar 2 cause adult type II citrullinemia, a rare disease marked by a sudden onset of disorientation, restlessness, drowsiness and coma, leading in the majority to death within a few years due to cerebral edema (Yasuda et al. 2001). GC1 (SLC25A22) and GC2 (SLC25A23) are glutamate proton symporters in the mitochondrial inner membrane (Fiermonte et al. 2001). They have chiefly been implicated in the regulation of amino acid deamination in association with glutamate dehydrogenase (LaNoue and Schoolwert 1979; Bradford and McGivan 1973). GC1 is widely distributed in human tissues with high levels in pancreas, liver, brain and testis and lower levels in heart, kidney, lung, small intestine and spleen (Fiermonte et al., 2001). In contrast, GC2 expression is largely confined to the brain and testis (Fiermonte et al. 2001). When the carriers were reconstituted into proteoliposomes their substrate specificity was virtually confined to glutamate; they were potently inhibited by -cyanocinnamate and bongkrekic acid; the Km for GC1 was around 5 mmoles, whilst that for GC2 was approximately 0.25 mmoles (Fiermonte et al. 2001). A recent report has suggested that a member of the excitatory amino acid transporter (EAAT) family, EAAT1 is expressed in cardiac mitochondria (Ralphe et al. 2004). This transporter family, which contains 5 members, constitutes the high affinity sodium dependent glutamate and aspartate transporters that are most easily recognizable for their contribution to neurotransmitter re-uptake in the central nervous system (Kanai and Hediger 2004). It is known that EAAT1EAAT3 are present in heart (King et al. 2004; Kugler 2004), where EAAT2 and EAAT3 are believed to reside within the cardiac sarcolemma (King et al. 2001; King et al. 2004; Kugler 2004). The exact function of EAAT1 in cardiac mitochondria remains to be determined, although Ralphe et al. (2005) proposed that it may be an additional aspartate-glutamate carrier involved in the malateaspartate shuttle.

6.2.3. Neutral Amino Acids Early investigations of substrate flux across the mitochondrial inner membrane suggested there was a single neutral amino acid carrier with a broad specificity (reviewed in LaNoue and Schoolwerth 1979; Porter 2000). At this time, proline was noted as a likely exception since it was not a very effective agent in protecting other neutral amino acids against mersalyl inhibition (LaNoue and Schoolwerth 1979), yet proline oxidation occurs exclusively in the mitochondria (Passarella et al. 2003). Subsequently, Atlante et al. (1994) demonstrated in rat kidney mitochondria the existence of a proline uniporter and a proline-glutamate antiporter (Figure 6.1), the latter of which was dependent upon the membrane potential difference.

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Another amino acid that has sparked controversy regarding its transport mechanism is glutamine. The discussions seem to have been prompted by investigations of the regulation of glutaminase activity, which catalyzes the deamination of glutamine to glutamate and ammonia in the matrix, and the extent to which this is limited by glutamine entry across the inner mitochondrial membrane (LaNoue and Schoolwerth 1979; Passarella et al. 2003). This dispute has yet to be resolved; however the purification and reconstitution into liposomes of a 41.5 kiloDalton carrier protein has helped clarify the specifics of glutamine transport (Indiveri et al. 1998). The reconstituted glutamine carrier operated either as a uniporter or as a homoexchanger (Indiveri et al. 1998). Glutamine uptake was highest when the mitochondria were solubilized at pH 6.5 and the protein was reconstituted at the same pH (Indiveri et al. 1998). The only other amino acids tested, amongst a group containing aspartate, glutamate, leucine, alanine, glycine, proline and serine, that was transported was asparagine (Indiveri et al. 1998).

6.2.4. Basic Amino Acid Carriers A basic amino acid transporter has been described in brain mitochondria (Porter 2000; Dolinska et al. 1998). The uptake of arginine was sensitive to lysine and ornithine suggesting that these compounds were also substrates with the uptake inhibited by the nitric oxide inhibitor L-N-monomethyl arginine, but neither glutamine nor histidine nor glutamate affected transport (Dolinska et al. 1998). The Km for arginine uptake in rat cerebral mitochondria was 0.08 mmoles (Dolinska et al. 1998).

6.3. Amino Acid Metabolism in the Mitochondria Under Normal Conditions 6.3.1. Introduction As important intermediary metabolites, amino acids are involved in numerous major pathways many of which have a mitochondrial component. These different reactions may contribute to: energy production, including for example gluconeogenesis, ketogenesis and supplementation of citric acid cycle intermediates (Jungas et al. 1992; Hutson et al. 2005; Pisarenko 1996); cellular and organ ionic homeostasis, such as the regulation of nitrogen (Hutson et al. 2005; Medina 2001) and sulfur levels (Townsend et al. 2004); and the synthesis of many biologically active compounds e.g. purines, pyramidines, hormones, neurotransmitters and other proteins (Mathews and Van Holde 1990). A comprehensive review of all these processes would be worthy of a complete volume and as such is beyond the scope of this chapter. Nonetheless, it would be neglectful to ignore all the metabolic pathways involving amino acids that occur in the mitochondria. We will therefore restrict ourselves to discussing just two of arguably the most significant pathways namely: the malate-aspartate shuttle and the urea cycle.

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6.3.2. The Malate-Aspartate Shuttle It has long been known that the mitochondrial inner membrane is impermeable to the NADH/NAD+ couple (Lehninger 1951). This presents a problem since the initial stages of energy production from glucose or lactate yield pyruvate and NADH in the cytoplasm. In order to achieve maximal ATP production this cytoplasmic pyruvate must be transferred into the mitochondria for oxidative metabolism within the citric acid cycle accompanied by an equivalent oxidation of NADH also within the mitochondria (Safer 1975). The existence of a “shuttle” utilising the cytoplasmic aspartate aminotransferase and the mitochondrial malate dehydrogenase (Figure 6.2), which could achieve redox segregation was initially proposed in 1963 (Borst). However, Borst (1963) was not entirely confident in his scheme, because the system was symmetrical resulting in at equilibrium equal NAD+ /NADH ratios in the cytoplasm and the mitochondria. Previous work had shown that the NAD+ /NADH ratio was more oxidized in the cytoplasm than the mitochondria (Williamson et al. 1967). Two further pieces of evidence were required before the malate-aspartate shuttle as shown in Figure 6.2 came to be recognised as a vital conduit of cellular metabolism.

Figure 6.2. The malate-aspartate shuttle. Arrows indicate the direction of reactions. Enzymes are numbered: 1 Cytoplasmic glutamate-oxaloacetate transaminase; 2 cytoplasmic malate dehydrogenase; 3 Mitochondrial glutamate-oxaloacetate transaminase; and 4 mitochondrial malate dehydrogenase. Aralar 1 or 2, the aspartate-glutamate exchanger; and OGC the 2-oxoglutarate (i.e. -ketoglutarate) carrier.

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The first clue was that in coupled mitochondria transfer of electrons in the respiratory chain results in a small pH gradient (inside alkaline) and a large electrical potential difference (inside negative) across the mitochondrial inner membrane (Safer 1975). The other related discovery was the observation that the exchange of the acidic amino acids aspartate and glutamate was mediated by a specific carrier in the inner membrane and was electrogenic in nature (LaNoue and Schoolwerth 1979). More specifically, a cotransported proton drove glutamate into the matrix, whilst an uncharged aspartate ion moved in the opposite direction (LaNoue 1974). The significance of this is that the other carrier involved in the shuttle and localised within the inner membrane, OGC (SLC20A4, Iacobazzi et al. 1992) mediates the electroneutral exchange of cytoplasmic malate for -ketoglutarate in the matrix (Kaplan 2001). Hence the aspartate-glutamate exchanger is widely regarded as the rate-limiting step in the operation of the shuttle (Palmieri et al. 2001).

6.3.3. The Urea Cycle Ammonia is produced as a by-product of protein and amino acid catabolism. It has been estimated that 505 mmoles of ammonia are generated and transferred to the liver each day (Jungas et al. 1992). Without further metabolism such levels would be toxic as exemplified by several disease states in which excess ammonia accumulates, resulting in for example brain edema, mental retardation and cerebral palsy (Butterworth 2001). Under normal conditions, this is prevented through a cyclical chain of reactions (Figure 6.3) wherein ammonia is converted by the liver into urea, which can then be excreted via the kidneys. The urea cycle utilizes the amino acids, aspartate, arginine, citrulline and ornithine. The only one of these to be either consumed or produced is aspartate, which is required for the reaction catalyzed by argininosuccinate synthetase (ASS). Some of this aspartate may originate from transamination of oxaloacetate in the matrix followed by transport on the glutamate-aspartate exchanger, aralar 2 that is an integral part of the malate-aspartate shuttle (Palmieri et al. 2001, see above and Figures 6.1 and 6.2). Recently, it has been shown that in the fetal liver 90% and in adult liver 30% of ASS expression is located on the outer mitochondrial membrane although most texts designate this reaction as occurring in the cytoplasm (Husson et al. 2003). The next amino acid, arginine is catabolised by arginase. In hepatocytes this reaction is mostly catalysed by a cytoplasmic enzyme, arginase I (Cheung and Raijman 1981). A second enzyme, arginase II, which is responsible for about 10% of the total hepatocyte arginase activity (Cheung and Raijman 1981), is present in the inner mitochondrial membrane. Arginase II has a wide tissue distribution (Colleluori et al. 2001). It has a minor role supplementing urea cycle activity in liver (Nissim et al. 2005). In addition it has been implicated in the biosynthesis of proline and polyamines (Wu and Morris 1998), and, through its influence on the intracellular arginine pool, as a regulator of NO2 availability and smooth muscle relaxation in opossum anal internal sphincter (Baggio et al. 1999).

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Figure 6.3. The urea cycle. Reactions involving the enzymes 1–3 occur in the cytoplasm, whilst reactions involving the enzymes 4–5 occur in the mitochondrial matrix. A heavy continuous line representing the mitochondrial inner membrane and a heavy dashed line representing the mitochondrial outer membrane indicate this segregation. Arrows indicate the direction of reactions. Enzymes are numbered: 1 Argininosuccinate synthetase; 2 Argininosuccinase; 3 Arginase; 4 Ornithine transcarbamoylase; and 5 Carbamoyl phosphate synthetase.

Finally, the only reactions that are definitively assigned to the mitochondrial matrix are those involving the enzymes: ornithine transcarbomoylase and carbomoyl phosphate synthetase (Jungas et al. 1992, Nissim et al. 2005), which in essence add free NH3 onto ornithine to yield citrulline.

6.4. Amino Acids in Mitochondria Under Pathological Conditions 6.4.1. Introduction In this section the role of amino acids in three different pathological states will be examined namely: ischemia-reperfusion injury, hypertrophy and cancer.

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For the sake of simplicity the first two conditions will be discussed as they pertain to cardiac mitochondria, whilst cancer will be discussed in relation to glutamine.

6.4.2. Ischemia-Reperfusion Injury When blood is diverted away from the heart (ischemia) this causes a decrease in the supply of nutrients and oxygen to the myocardium. This leads to a reduction in energy production in the mitochondria, a fall in intracellular pH, an increase in [Ca2+ ]i and [Na+ ]i and an accumulation of abnormal metabolites e.g. lactic acid (reviewed in Suleiman et al. 2001). If the ischemic period is short, restoration of the blood flow (reperfusion) enables cellular recovery; however after prolonged ischemia, reperfusion may result in significant cellular and tissue damage. The foundation to the proposal that amino acids may alleviate ischemiareperfusion injury was largely based on their potential to convert to citric acid cycle intermediates with subsequent substrate level phosphorylation and ATP production in the mitochondria (Pisarenko 1996; Taegtmeyer 1995; Sveldjeholm et al. 1995). Logical, though this may be, the concept does not enjoy universal support (Buckberg 1996). A likely cause of this controversy is the relatively poor understanding of the mechanisms of amino acid transport across the cardiac sarcolemma. Thus, in the majority of cases where the addition of exogenous amino acids leads to an increase in their myocardial concentration the outcome has been beneficial (Pisarenko et al. 1985; Pisarenko et al. 1995). Branched chain amino acids are transported across the cardiac sarcolemma by the L system of mediation (King et al. 2001). The initial stages of their metabolism occur exclusively in the mitochondria where the branched chain aminotransferase is located (Hutson et al. 1988). Schwalb et al. (1989) exposed working rat hearts to cardioplegic arrest followed by 68 min of global normothermic ischemia and 40 min reperfusion. The control group received a modified St Thomas solution whilst the test groups received the same solution with 11.1 mmoles glucose plus or minus 3.5 mmoles branched chain amino acids (BCAA) (Schwalb et al. 1989). During reperfusion ATP levels and the recovery of cardiac output were significantly higher in the group receiving BCAA than in either of the other two groups (Schwalb et al. 1989). Similarly, addition of acidic amino acids to crystalloid or blood cardioplegia has also been shown to improve functional recovery and ATP levels in various animal hearts (Haas et al., 1984; Lazar et al. 1980; Bittl and Shine 1983; King et. al. 2004). Oxidative stress accompanied by the production of reactive oxygen species (ROS) is one of the major pathogenesis of ischemia-reperfusion (Suleiman et al. 2001; Zweier et al. 1988). Figure 6.4 illustrates the results of experiments carried out using freshly isolated adult rat cardiomyocytes, which show that complex III of the respiratory chain in the mitochondria can be a major source of these ROS. This experiment was performed using the inhibitors, antimycin A, which prolongs the lifespan of ubisemiquinone thereby promoting ROS production, and myxothiazol, which prevents electron transfer to ubisemiquinone consequently

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Figure 6.4. Mitochondrial production of reactive oxygen species in freshly isolated rat ventricular cardiomyocytes. The fluorescence of the probe, 5-(and-6)-chloromethyl2’, 7’-dichlorodihydrofluorescein diacetate (CM-DCFda) is proportional to the level of hydroxy radical generation. This graph shows a time course of the fluorescence measured in freshly isolated rat ventricular cardiomyocytes that have been loaded with 5 moles CM-DCFda then superfused with a control Tyrode solution or a Tyrode solution containing either 10 moles myxothiazol or 1 mole antimycin A. The increase in fluorescence in the presence of antimycin A and the accompanying lack of effect of myxothiazol suggest that complex III of the mitochondrial respiratory chain can be a source of hydroxy radicals in cardiomyocytes. *p < 0.01 vs. control and + myxothiazol. Results shown are the means ±S.D (n = 4).

lowering ROS generation (King et al. 2003). When cardiomyocytes are isolated in the presence of glutamate this results in a higher NAD+ /NADH ratio, higher ATP levels (Williams et al. 2001) and lower levels of ROS production during exposure to antimycin A or metabolic inhibition compared to non-glutamate loaded controls (King et al. 2003). Although the exact mechanism behind the antioxidant effect of glutamate remains undetermined, it has been speculated that a pathway may exist linking the activity of glutamate dehydrogenase in the mitochondria through glutathione reductase to glutathione peroxidase and the detoxification of H2 O2 to H2 O (Williams et al. 2001) (Figure 6.5).

6.4.3. Hypertrophy Myocardial hypertrophy is an adaptive response to a chronic increase in the heart’s workload. It occurs in several pathophysiological conditions including sustained hypertension, aortic constriction, valvular insufficiency and hyperthyroidism (Rossi and Lortet 1996). Its onset and development is associated with morphological and physiological restructuring of the heart that will affect all

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Glutamate

α-Ketoglutarate 1 +

NADPH

NADP

2 2 GSH

GSSG

3 H2O

H2O2

Figure 6.5. Hypothetical scheme linking glutamate metabolism in the mitochondria to the detoxification of hydrogen peroxide. Glutamate deamination in the mitochondria through 1: glutamate dehydrogenase using the NADP+ /NADPH couple is linked through the activities of 2: glutathione reductase and 3: glutathione peroxidase to the detoxification of hydrogen peroxide to water. Arrows indicate the directions of reactions. 2 GSH reduced glutathione, GSSG oxidized glutathione.

of the organelles and cellular functions (Rossi and Lortet 1996). The following discusses one of the ways in which these changes influence amino acids in the cardiac mitochondria. In several models, the progression of cardiac hypertrophy is marked by increased glucose transport through GLUT1 (Montessuit and Thorburn 1999), higher activity levels of the enzymes involved in glycolysis (Seymour et al. 1990; Valadares et al. 1969) and a down-regulation of fatty acid metabolism (El Alaoui-Talibi and Moravec 1989). This shift towards a more anaerobic pattern of ATP production might also be expected to impact upon the activity of the malate-aspartate shuttle. This indeed appears to be the case in hypertrophy induced by chronic exposure to the thyroid hormone, triiodothyronine (T3 ). In these hearts the capacity of the malate-aspartate shuttle is significantly increased by 33% compared to control (Scholz et al., 2000). This is accompanied in the T3 treated hearts by a significantly increased expression of the aspartate-glutamate carrier and mitochondrial malate dehydrogenase in comparison to control (Scholz et al. 2000). A more recent study by the same group, which benefited from the molecular identification of the glutamate-aspartate carriers (Palmieri et al. 2001), suggested that the acidic amino acid carrier whose expression was upregulated in hypertrophy was in fact EAAT1 (Ralphe et al. 2005). This supports work carried out in a model of chronic hypertension, the Spontaneously Hypertensive Rat, where increased EAAT expression was shown in hypertrophic hearts compared to control (King et al., 2004). The last mentioned investigation also showed that the addition of 0.5 mmoles aspartate to isolated and perfused hypertrophic hearts led to a significantly improved functional recovery upon reperfusion and higher ATP levels at the end of ischemia compared to hypertrophic hearts not receiving aspartate (King et al. 2004).

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6.4.4. Glutamine and Cancer A curious feature of tumor cells, as opposed to their normal counterparts, is their appetite for the neutral amino acid, glutamine (reviewed in Medina 2001 and Souba 1993). Rapidly growing cancer cells exhibiting high rates of nucleotide and protein synthesis require a continuous source of nitrogen (Medina 2001). Glutamine, as the most abundant amino acid in the body, is the main vehicle for circulating ammonia in a non-toxic form and is therefore a handy conduit for enabling this supply (Medina et al. 1992). Thereafter, the nitrogen can be liberated through the actions of glutaminase. This enzyme is located exclusively in the mitochondria where it catalyzes the conversion of glutamine to glutamate and NH3 (Queseda et al. 1988). There is a good correlation between the activity of glutaminase and the extent of malignant proliferation in various rat hepatomas (Linder-Horowitz et al. 1969) and breast cancer cells (Gómez-Fabre et al. 2000). Glutamine may also be the major respiratory fuel in tumor cells. In order to maintain optimum growth rates in conditions where the oxygen tension is variable and often low (Griffiths et al. 2001), tumor cells rely more heavily on glycolysis than on oxidative phosphorylation for ATP synthesis (Rossignol et al., 2004). This adaptation is characterized by a shift away from the expression of pyruvate kinase type L to the M2 type (Hacker et al. 1998, Ibsen, 1977). If glucose levels are very reduced a further metabolic revision occurs leading to the catabolism of glutamine for energy. Some studies go one step further and suggest that tumor cells can survive and grow in the absence of glucose as long as glutamine is present (Reitzer et al. 1979; Rossignol et al. 2004; Eagle et al. 1958). The pathway of glutamine catabolism in tumor cells is referred to as glutaminolysis (Figure 6.6) by analogy to glycolysis (McKeehan 1982). The second step in this pathway is a transamination reaction catalyzed by mitochondrial glutamate-oxaloacetate transaminase. This is more favorable than metabolism through glutamate dehydrogenase, because the high levels of NADH and lactate found in tumor cells tend to force this enzyme towards glutamate synthesis (Eigenbrodt et al. 1998). For example, malignant transformation of the oval cell line OC/CDE 22 resulted in decreased cytoplasmic glutamateoxaloacetate transaminase activity with concomitantly increased mitochondrial glutamate-oxaloacetate transaminase activity and no change in the activity of glutamate dehydrogenase compared to control cells (Mazurek et al. 1999). The final step in the glutaminolysis pathway involves oxidative decarboxylation catalyzed by the NAD+ dependent mitochondrial malic enzyme to yield pyruvate and CO2 (Loeber et al. 1991). This importance of glutamine to tumor cell metabolism and proliferation led to the attempted development in the 1980s of glutamine related anticancer therapies (Medina 2001). These approaches largely involved the use of amino acid analogs to reduce glutamine availability. Clinical trials were not however successful due to toxic effects, lack of specificity and/or ineffectiveness (Medina et al. 1992; Souba et al. 1993). More recently it has been shown that glutamine starvation leads to apoptosis of murine hybridoma cells following activation of

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Figure 6.6. Glutaminolysis. Schematic pathway showing the possible production of pyruvate from glutamine in the mitochondria of cancer cells. Arrows indicate the direction of reactions. Enzymes are numbered: 1 glutaminase; 2 mitochondrial glutamateoxaloacetate transaminase; 3 -ketoglutarate dehydrogenase; 4 succinyl-CoA synthetase; 5 succinate dehydrogenase; 6 fumarase and 7 malic enzyme.

the caspases –9 and –3 (Paquette et al. 2005). Perhaps as our understanding of the complexities of cellular signaling and the pathways of apoptosis (reviewed elsewhere in this volume) continue to advance this may open up new avenues for a glutamine-based anticancer therapy.

References Atlante A, Passarella S, Pierro P, Quagliariello E (1994) Proline transport in rat kidney mitochondria. Arch Biochem Biophys 309:139–148 Baggio R, Emig FA, Christianson DW, Ash DE, Chakdar S, Rattan S (1999) Biochemical and functional profile of a newly developed potent and isozyme-selective arginase inhibitor. J Pharmacol Exp Therapeut 290:1409–1416 Bittl JA, Shine KI (1983) Protection of ischemic rabbit myocardium by glutamic acid. Am J Physiol 245:H406–H412

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Borst P (1963) Hydrogen transport and transport metabolisms. In Karlson, P. (ed.), Functionelle und Morphologische Organisation der Zelle, Springer-Verlag, Berlin, pp. 137–158 Bradford NM, McGivan JD (1973) Quantitative aspects of glutamate transport in rat liver mitochondria. Biochem J 134:1023–1029 Buckberg GD (1996) Invited editorial on “Effects of glutamate and aspartate on myocardial substrate oxidation during potassium arrest”. J Thorac Cardiovasc Surg 112:1661–1663. Butterworth RF (2001) Glutamate transporter and receptor function in disorders of ammonia metabolism. Ment Retard Dev Disabil Res Rev 7:276–279 Cheung CW, Raijman L (1981) Arginine, mitochondrial arginase, and the control of carbamyl phosphate synthesis. Arch Biochem Biophys 209:643–649 Colleluori DM, Morris SM Jr, Ash DE (2001) Expression purification and characterization of human type II arginase. Archiv Biochem Biophys 389:135–143 Dolinska M, Albrecht J (1998) L-Arginine uptake in rat cerebral mitochondria. Neurochem Int 33:233–236 Eagle H, Barban S, Levy M, Schulz HO (1958) Utilization of carbohydrates by human cell cultures. J Biol Chem 233:551–558 Eigenbrodt E, Kallinowski F, Ott M, Mazurek S, Vaupel P (1998) Pyruvate kinase and the interaction of amino acid and carbohydrate metabolism in solid tumors. Anticancer Res 18:3267–3274 El Alaoui-Talibi Z, Moravec J (1989) Limitation of long chain fatty acid oxidation in volume over-loaded rat hearts. Adv Exp Med Biol 248:491–497 Fiermonte G, Palmieri L, Todisco S, Agrimi G, Palmieri F, Walker JE (2002) Identification of the mitochondrial glutamate transporter. J Biol Chem 277: 19289–19294 Gómez-Fabre PM, Aledo JC, Del Castillo-Olivares A, Alonso FJ, Núñezde Castro I, Campos JA, Márquez J (2000) Molecular cloning, sequencing and expression studies of the human breast cancer cell glutaminase. Biochem J 345:365–375 Griffiths JR (2001) Causes and consequences of hypoxia and acidity in tumor microenvironments. Bioessays 23:295–296 Haas GS, DeBoer LWV, O’Keefe DD, Bodenhamer RM, Geffin MB, Drop LJ, Teplick RS, Daggett WM (1984) Reduction of postischemic myocardial dysfunction by substrate repletion during reperfusion. Circ 70(suppl I):I-65–I73 Hacker HJ, Steinberg P, Bannasch P (1998) Pyruvate kinase isoenzyme shift from L-type to M2-type is a late event in hepatocarcinogenesis induced in rats by a cholinedeficient/DL-ethionine-supplemented diet. Carcinogenesis 19:99–107 Husson A, Brasse-Lagnel C, Fairand A, Renouf S, Lavoinne A (2003) Argininosuccinate synthetase from the urea cycle to the citrulline-NO cycle. Eur J Biochem 270: 1887–1899 Hutson SM, Fenstermacher D, Mahar C (1988) Role of mitochondrial transamination in branched chain amino acid metabolism. J Biol Chem 263:3618–3625 Hutson SM, Sweatt AJ, LaNoue KF (2005) Branched-chain amino acid metabolism: Implications for establishing safe intakes. J Nutr 135:1557S–1564S Iacobazzi V, Palmieri F, Runswick MJ, Walker JE (1992) Sequences of the human and bovine genes for the mitochondrial 2-oxoglutarate gene. DNA Seq 3:79–88 Ibsen KH (1977) Interrelationships and functions of the pyruvate kinase isoenzymes and their variant forms: a review. Cancer Res 37:341–353 Indiveri C, Abruzzo G, Stipani I, Palmieri F (1998) Identification and purification of the reconstitutively active glutamine carrier from rat kidney mitochondria. Biochem J 333:285–290

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Jungas RL, Halperin ML, Brosnan JT (1992) Quantitative analysis of amino acid oxidation and related gluconeogenesis in humans. Physiol Rev 72:419–448 Kanai Y, Hediger MA (2004) The glutamate/neutral amino acid transporter family SLC1: molecular, physiological and pharmacological aspects. Pflügers Archiv 447:469–479 Kaplan RS (2001) Structure and function of mitochondrial anion transport proteins. J Membrane Biol 179:165–183 King N, Lin H, McGivan JD, Suleiman MS (2004) Aspartate transporter activity in hypertrophic rat heart and ischaemia-reperfusion injury. J Physiol 556: 849–858 King N, McGivan JD, Griffiths EJ, Halestrap AP, Suleiman MS (2003) Glutamate loading protects freshly isolated and perfused adult rat cardiomyocytes from intracellular ROS generation. J Mol Cell Cardiol 35: 975–984 King N, Suleiman MS (2001) L-Leucine transport in rat heart under normal conditions and effect of simulated hypoxia. Mol Cell Biochem 221: 99–108 King N, Williams H, McGivan JD, Suleiman MS (2001) Characteristics of L-aspartate transport and expression of EAAC-1 in sarcolemmal vesicles and isolated cells from rat heart. Cardiovasc Res 52: 84–94 Kugler P (2004) Expression of glutamate transporters in rat cardiomyocytes and their localization in the T-tubular system. J Histochem. Cytochem 52:1385–1392 LaNoue KF, Tischler ME (1974) Electrogenic characteristics of the mitochondrial glutamate-aspartate antiporter. J Biol Chem 249:7522–7528 LaNoue KF, Schoolwerth AC (1979) Metabolite transport in mitochondria. Ann Rev Biochem 48:871–922 Lazar HL, Buckberg GD, Manganaro AM, Becker H (1980) Myocardial energy replenishment and reversal of ischemic damage by substrate enhancement of secondary blood cardioplegia with amino acids during reperfusion. J Thorac Cardiovasc Surg 80:350–359 Lehninger AL (1951) Phosphorylation coupled to oxidation of dihydrodi-phosphopyridine nucleotide. J Biol Chem 190:345–359 Linder-Horowitz M, Knox WE, Morris HP (1969) Glutaminase activities and growth rates of rat hepatomas. Cancer Res 29:1195–1199 Mathews CK, Van Holde KE (1990) Biochemistry. Benjamin Cummings Publishing Company, Redwood City, California McKeehan WL (1982) Glycolysis, glutaminolysis and cell-proliferation. Cell Biol Int Rep 6: 635–650 Medina MA (2001) Glutamine and cancer. J Nutr 131:2539S–2542S Medina MA, Sánchez-Jiménez F, Márquez J, Queseda AR, Núñez de Castro I (1992) Relevance of glutamine metabolism to tumor cell growth. Mol Cell Biochem 113:1–15 Montessuit C, Thorburn A (1999) Transcriptional activation of the glucose transporter GLUT1 in ventricular cardiomyocytes by hypertrophic agonists. J Biol Chem 274: 9006–9012 Nissim I, Luhovyy B, Horyn O, Daihin Y, Nissim I, Yudkoff M (2005) The role of mitochondrially bound arginase in the regulation of urea synthesis. J Biol Chem 280:17715–17724 Palmieri L, Pardo B, Lasorsa FM, Del Arco A, Kobayashi K, Lijima M, Runswick MJ, Walker JE, Saheki T, Satrústegui J, Palmieri F (2001) Citrin and aralar1 are Ca2+ -stimulated aspartate/glutamate transporters in mitochondria. EMBO J 20: 5060–5069 Paquette JC, Guérin PJ, Gauthier ER (2004) Rapid induction of the intrinsic apoptotic pathway by glutamine starvation. J Cell Physiol 202:912–921 Passarella S, Atlanate A, Valenti D, De Bari L (2003) The role of mitochondrial transport in energy metabolism. Mitochondrion 2:319–343

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Pisarenko OI (1996) Mechanisms of myocardial protection by amino acids: facts and hypotheses. Clin Exptl Pharmacol 23:627–633 Pisarenko OI, Novikova EB, Serebryakova LI, Tskitishvili OV, Ivanov VE, Studneva IM (1985) Function and metabolism of dog heart in ischaemia and in subsequent reperfusion: effect of exogenous glutamic acid. Pflügers Archiv 405:377–383 Pisarenko OI, Studneva IM, Shulzhenko VS, Korchazhkina OV, Kapelko VI (1995) Substrate accessibility to cytosolic aspartate aminotransferase improves posthypoxic recovery of isolated rat heart. Biochemical Mol Med 55:138–148 Porter RK (2000) Mammalian mitochondrial inner membrane cationic and neutral amino acid carriers. Biochim Biophys Acta 1459:356–362 Queseda AR, Sánchez-Jiménez F, Perez-Rodriguez J, Márquez J, Medina MA, Núñez de Castro I (1988) Purification of phosphate-dependent glutaminase from isolated mitochondria of Ehrlich ascites tumor cells. Biochem J 255:1031–1036 Ralphe JC, Bedell K, Segar JL, Scholz TD (2005) Correlation between myocardial malate/aspartate shuttle activity and EAAT1 protein expression in hyper- and hypothyroidism. Am J Physiol. 288:H2521–H2526 Ralphe JC, Segar JL, Schutte BC, Scholz TD (2004) Localization and function of the brain excitatory amino acid transporter type 1 in cardiac mitochondria. J Mol Cell Cardiol 37:33–41 Reitzer LJ, Wice BM, Kennell D (1979) Evidence that glutamine, not glucose, is the main energy-source for Hela-cells. J Biol Chem 254:2669–2676 Rossi A, Lortet S (1996) Energy metabolism patterns in mammalian myocardium adapted to chronic pathophysiological conditions. Cardiovasc Res 31:163–171 Rossignol R, Gilkerson R, Aggeler R, Yamagata K, Remington SJ, Capaldi RA (2004) Energy substrate modulates mitochondrial structure and oxidative capacity in cancer cells. Cancer Res 64: 985–993 Safer B (1975) The metabolic significance of the malate-aspartate cycle in heart. Circ Res 37, 527–533 Scholz TD, TenEyck CJ, Schutte BC (2000) Thyroid hormone regulation of the NADH shuttles in liver and cardiac mitochondria. J Mol Cell Cardiol 32:1–10 Schwalb H, Yaroslavsky E, Borman JB, Uretzky G (1989) The effect of amino acids on the ischemic heart. J Thorac Cardiovasc Surg 98:551–556 Seymour AML, Eldar H, Radda GK (1990) Hyperthyroidism results in increased glycolytic capacity in the rat heart. A 31 P-NMR study. Biochim Biophys Acta 1055:107–116 Souba WW (1993) Glutamine and cancer. Ann. Surg 218:715–728 Suleiman MS, Halestrap AP, Griffiths EJ (2001) Mitochondria: a target for myocardial protection. Pharmacol Therapeut 89:29–46 Sveldjeholm R, Håkanson E, Vanhanen I (1995) Rationale for metabolic support with amino acids and glucose-insulin-potassium (GIK) in cardiac surgery. Ann Thorac Surg 59:S15–S22 Taegtmeyer H (1995) Metabolic support for the postischaemic heart. Lancet 345: 1552–1555 Townsend DM, Tew KD, Tapiero H (2004) Sulfur containing amino acids and human disease. Biomed Pharmacother 58:47–55 Valdares JRE, Singhai RL, Parulekar MR, Beznak M (1969) Influence of aortic coarctation on myocardial glucose-6-phosphate dehydrogenase. Can J Physiol Pharmacol47: 388–391 Williams H, King N, Griffiths EJ, Suleiman MS (2001) Glutamate-loading stimulates metabolic flux and improves cell recovery following chemical hypoxia in isolated cardiomyocytes. J Mol Cell Cardiol 33: 2109–2119

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Williamson DH, Lund P, Krebs HA (1967) Redox state of free nicotinamide-adenine dinucleotide in cytoplasm and mitochondria of rat liver. Biochem J 103:514–527 Wu GY, Morris SM (1998) Arginine metabolism: nitric oxide and beyond. Biochem J 336:1–17 Yasuda T, Yamaguchi N, Kobayashi K, Nishi I, Horinouchi H, Jalil A, Li MX, Ushikai M, Lijima M, Kondo I, Saheki T (2000) Identification of two novel mutations in the SLC25A13 gene and detection of seven mutations in 102 patients with adult-onset type II citrullinaemia. Hum Genet 107:537–545 Zweier JL (1988) Measurement of superoxide-derived free radicals in the reperfused heart. J Biol Chem 263:1353–1357

Part 2 The Dynamic Nature of the Mitochondria

7 Mechanotransduction of Shear-Stress at the Mitochondria Abu-Bakr Al-Mehdi

7.1.

Mechanotransduction of Shear–Stress

7.1.1. Physical Forces due to Blood Flow Endothelial cells in situ experience a variety of physical forces caused by hemodynamics. Tensile stress (a perpendicular force of pressure and stretch) affects mostly the smooth muscle cells, and shear-stress (a tangential, frictional force) affects specifically the endothelial cells. Endothelial cells that are flow-adapted in vivo or in flow-chambers are different from their statically grown counterparts. However, the common practice of growing endothelial cells in Petri dishes initially overlooked this natural environmental physical factor until its role was convincingly recognized in endothelial cell physiology in the early eighties (Dewey et al. 1981). These physical factors and their role in endothelial cell function have drawn increased attention recently (Lansman 1988; Davies and Tripathi 1993). Hemodynamic forces generated by the flow of blood along the endothelial surface are now known to regulate both the morphology and function of endothelial cells in different physiological and patho-physiological settings. Endothelial cells transform the mechanical stimuli into electrical and biochemical signals in a process called mechano-transduction. Although the precise mechanisms of cellular mechanotransduction have yet to be elucidated, tensile and shear-stress responsive ion channels and cytoskeletal elements may be involved. Once the role of physical forces in endothelial cell biology was recognized, researchers started studying the effects of shear-stress by applying flow to statically cultured endothelial cells in a variety of devices, like cone viscometers, parallel plate chambers, and artificial capillary tubes. These studies of the effect of increased laminar, turbulent and oscillatory flow on endothelial cell function have relevance for elucidating the pathogenetic mechanisms of hypertension, arteriosclerosis, and other intravascular disorders. Time-dependent responses of endothelial cells subjected to laminar shear-stress 169

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include: (1) rapid (within 1 min) hyperpolarization of the cell membrane and activation of G proteins, (2) a slower (1–60 min) activation of MAP kinases and NF-B, cytoskeletal rearrangement, induction of connexin 43 and cell adhesion proteins, (3) realignment of focal adhesions by 1–6 h, and (4) a delayed (6–24 h) reorganization of cell surface and cellular alignment in the direction of flow (Davies and Tripathi, 1993). After 6 to 24 hours of shear-stress exposure, these cells can be considered ‘flow adapted’ analogous to their natural counterparts in vivo. Strikingly, some responses of the endothelial cells were shown to be similar for both types of stimuli, i.e., increased shear-stress in non-adapted cells or loss of shear-stress in adapted cells. These responses include pinocytosis (Davies et al. 1984), intracellular calcium (James et al. 1995; Tozawa et al. 1999), reactive oxygen species generation (Hsieh et al. 1998; Wei et al. 1999), and nitric oxide generation (Mashour and Boock 1999; Al-Mehdi et al. 2000a). That is, the response of flow adapted endothelial cells to loss of shear-stress is similar in direction to the response following onset of flow for cells adapted to static conditions. These responses represent a paradigm of alteration from a state of adaptation (Davies 1995), that is, homeostasis at a new baseline level after the subsidence of acute responses to flow or no-flow. The acute responses themselves are the result of sensing increased shear-stress when flow is instituted over endothelial cells adapted to no-flow; or they represent its inverse, the result of sensing loss of shear-stress when flow is stopped for endothelial cells adapted to shear-stress, as would happen during ischemia in vessels in vivo. The high degree of adaptability of endothelial cells allows them to cope with the challenges of hemodynamic alterations that they are constantly exposed to even under physiological conditions and thereby contributing to their fitness for survival. It is theorized that the sensing of shear-stress proceeds via specialized sensors, which may be specific plasma membrane molecules and microdomains, or via alterations in general biophysical forces such as membrane fluidity and cytoskeletal tension (Davies et al. 2003). The forces of shear-stress may act locally as the sensors detect them, or they may be transmitted to distant sites mechanically by means of the cytoskeleton, to elicit responses from organellar, nuclear, cell-cell junction, and focal adhesion mechanotransducers. The cis-acting DNA sequences that confer shear-mediated transcriptional activation are not well characterized. Two potential candidates for such a shearstress response element have been suggested. The first is the SSRE (shear-stress response element) identified as a 6-bp consensus sequence GAGACC, first described for the gene encoding platelet-derived growth factor B (Resnick and Gimbrone 1995). The second is a CT-rich Sp1 binding site within the vascular endothelial growth factor receptor (i.e., FIk-1/KDR) that showed increased occupancy after 3 hours of fluid shear-stress (Abumiya et al. 2002). It is known that shear-stress affects nuclear gene expression; hundreds of genes are up- or down-regulated under a variety of shear-stress conditions (Chen et al. 2001; Garcia-Cardena et al. 2001; McCormick et al. 2001; Brooks et al.

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2002; Kallmann et al. 2002; Peters et al. 2002). It is not known however, if shear-stress affects the expression of mitochondrial genes.

7.1.2. The Mechanotransduction Hypothesis of Endothelial Cell Response to Ischemia Ischemia or termination of blood flow in organs with systemic circulation not only results in loss of oxygen, reduced substrate supply, and accumulation of metabolites, but also the loss of shear-stress for the endothelial cells. The models of hypoxia/reoxygenation performed on statically grown endothelial cells do not take into account the effects caused by the lack of shear-stress. In the lung, occlusion of the pulmonary artery in the presence of ventilation (as in the case of clinical pulmonary thromboembolism) would not lead to tissue anoxia, but would result in the loss of shear-stress. In fact, during ischemia, local lung tissue pO2 will rise because oxygen is no longer removed by capillary blood. Therefore, an isolated, air-ventilated, artificial-medium perfused rat or mouse lung not only serves as an excellent model to study endothelial cells in situ, but also allows the effects of anoxia, non-hypoxic ischemia, and the loss of shear to be separated. Our experience with this isolated lung model and the development of an intact organ fluorescence imaging technique to study changes in endothelial cell function in situ led credence to the mechanotransduction hypothesis of ischemia (Al-Mehdi et al. 1997a; Al-Mehdi et al. 1998a; Al-Mehdi et al. 1998b). This hypothesis provides an explanation for the generation of reactive oxygen species (ROS) by the flow-deprived, but non-hypoxic endothelial cells, (Al-Mehdi et al. 1994; Al-Mehdi et al. 1996; Al-Mehdi et al. 1997a; AlMehdi et al. 1997b; Al-Mehdi et al. 1998a; Al-Mehdi et al. 1998b; Wei et al. 1999; Al-Mehdi et al. 2000a; Al-Mehdi et al. 2000b; Song et al. 2001; Wei et al. 2001; Fisher et al. 2002; Fisher et al. 2003; Wei et al. 2004). The basic premise of the hypothesis is that ischemia is sensed by the endothelial cells of the non-hypoxic lung as a loss of shear-stress, an inverse phenomenon of sensing the imposition of shear-stress by statically cultured endothelial cells upon resumption of flow. Therefore, the ischemic response of endothelial cells is a mechanotransduction event. Recent data suggest a direct role of the mitochondria in flow-sensing and the responses to shear-stress. We postulate that the mitochondrial response is more important for the long-term effects of flow or no-flow, such as cellular and organellar adaptation. Apparently, the endothelial cells must acquire the ability to respond to the loss of shear-stress during flow-adaptation.

7.2.

Mitochondrial Mechanotransduction

7.2.1. Why can Mitochondria be Mechanotransducers? The frictional force that is produced by blood flow is localized at the plasma membrane. It is readily apparent that this force can act locally on

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membrane-associated mechanosensors. Membrane proteins such as integrins, G-proteins, ion channels and localized rigid domains (cholesterol rich caveolae and lipid rafts) can serve as mechanosensors. As opposed to the localized sensing via molecular or domain-specific mechanisms, global sensing mechanisms are thought to be mediated via alterations in plasma membrane viscosity (or fluidity) and cytoskeletal tension (‘tensegrity’ as in Stamenovic 2005). The mechanical force of shear-stress can be transmitted to non-apical parts of the cell via cytoskeletal elements. The cytoskeleton has been shown to transmit the mechanical force of shear-stress to intracellular sites. Indeed, the alignment of actin filaments of macrovascular endothelial cells in the direction of flow is responsible for overall cellular alignment and elongation (Malek and Izumo 1996). Association of mitochondria with the cytoskeleton makes them potential sites of mechanotransduction. Mitochondrial movement uses the molecular motors kinesin and dynein that associate with microtubules and tether mitochondria to them. Microtubules are the thickest and most rigid of all cytoskeletal fibers; therefore, they are more suitable for transmission of mechanical force. The transmitted force of shear-stress, acting on points of contact between microtubules and organelles or basolateral focal adhesion sites, may be transduced locally to produce a signaling event. Mechanotransmission via microtubules, but not via intermediate or microfilaments, has been implicated in nuclear morphology changes with shear-stress (Stamatas and McIntire 2001). The mechanotransduction of shear-stress can be modeled analogous to ligandreceptor signal transduction. For flow-mediated signaling, the ‘ligand’ or primary signal is the mechanical force and the ‘receptor’ of this force is a mechanotransducer. The downstream components of mechanotransduction signaling share the same second messengers and biochemical pathways as the conventional ligandreceptor systems. Once the mechanical force has been transmitted to the mitochondrial membrane, a mechanotransducer on the mitochondrial membrane (inner or outer) would be required to convert the force into a biochemical signal. The identity of mitochondrial mechanotransducers is less clear than the mechanotransducers of the plasma membrane. The mtKATP channel, the Na+ /H+ , Na+ /Ca2+ , and K+ /H+ exchangers, and the mitochondrial permeability transition pore can be considered as potential mechanotransducers. Stretch, a mechanical force that is experienced by vascular cells upon distension, has been shown to affect cellular responses with mitochondrial participation (Ali et al. 2004; Liao et al. 2004). Actin filaments are the other cytoskeletal elements that need to be considered for mechanotransmission. If microtubules are tracks for movement, then actin filaments are stations for mitochondrial clustering or stopping (Boldogh and Pon 2006). Although actin filaments have lower elasticity than microtubules, their bundled and networked structure may be strong enough to transmit mechanical forces from the plasma membrane to the stationary mitochondria. The actin network has been shown to significantly contribute to cellular rigidity (Janmey et al. 1991).

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7.2.2. Effect of Flow and Flow Cessation on Mitochondrial Function Exposure of endothelial cells to flow or ischemia leads to changes in mitochondrial shape, size, position, number, and functional status. A shear-stress of 10 dyn/cm2 increases mitochondrial membrane potential by 30%; but a higher shear-stress (60 dyn/cm2 ) decreases mitochondrial membrane potential by 20% (Kudo et al. 2000). Mitochondria of lung pulmonary microvascular endothelial cells subjected to 1–10 dynes/cm2 laminar flow rapidly hyperpolarize (increased JC-1 fluorescence) upon cessation of flow, suggesting that the mitochondria sense and functionally respond to the loss of shear-stress (Figure 7.1). The response is seen in both pulmonary macro- and microvascular cell types after flow-adaptation. Acquisition of the hyperpolarization response to flow-cessation in flow-adapted macro- and macrovascular cell types is a paradigm of common functional adaptation. In contrast, only pulmonary macrovascular cells seem to exhibit signs of morphological adaptation to flow by elongation and alignment in the direction of the flow. This dissociation of morphological and functional adaptive responses to flow in the two cell types suggests the presence of separate mechanisms of flow-mediated signal transduction. Microtubule disruption with 1 μM colchicine prevents flow cessation-induced mitochondrial hyperpolarization, suggesting that microtubules are involved in mechanotransmission (Figure 7.2). Moreover, the depolarization response to flow-cessation suggests that mitochondrial membrane potential is coupled to shear-stress via some other mechanisms. A role for the actin cytoskeleton may explain this ‘paradoxical’ depolarization response to the loss of shear-stress. A microtubular role for maintenance of mitochondrial membrane potential is

Control

Flow-cessation

Figure 7.1. The left panel shows intensity plot of a rat pulmonary artery endothelial cell flow-adapted for 20 hours at 10 dynes/cm2 and stained with JC-1 (Control). Right panel (Flow-cessation) shows the same cell after flow was stopped for 1 min while maintaining oxygenation. The increase in JC-1 red aggregate formation (indicated by increase in TRITC-channel fluorescence) indicates mitochondrial hyperpolarization. A purple-to white lookup table applied for pseudocolor rendering of intensity. Increases in green, yellow and red colors indicate that mitochondrial membrane hyperpolarization accompanies the loss of shear-stress.

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Figure 7.2. Flow-adapted endothelial cells were treated with 1 μM colchicine for 30 min before being subjected to flow-cessation. Panel on the left shows intensity plots of JC-1 aggregate fluorescence during flow showing polarized mitochondria. Flow-cessation in the presence of colchicine not only prevented mitochondrial hyperpolarization, but also led to a depolarization response. Table 7.1. Changes in  m with microtubule stabilization and disruption Condition

Control Colchicine Paclitaxel

JC-1 Aggregate Fluorescence, Arbitrary Units 1376 ± 89 (n=4) 756 ± 101 (n=3)* 2215 ± 176 (n=3)#

JC-1 Monomer Fluorescence, Arbitrary Units 768 ± 56 (n=4) 1024 ± 65 (n=3)* 460 ± 37 (n=3)#

Note: Mitochondria were imaged with a JC-1 filter set. The ratio of aggregate to monomer fluorescence, which increases at more negative  m , was 1.79 for control vs 0.73 and 4.8 upon colchicine and paclitaxel treatment, respectively. *p < 0.05 vs control; #p < 0.5 vs control and colchicine.

indicated by the contrasting effects of colchicine and paclitaxel (a microtubule stabilizer) (Table 7.1).

7.2.3. Mitochondrial Morphology and Function Under Shear-stress It has been demonstrated using MitoTracker Green that endothelial cells at branch points of lung capillaries contain two-fold more mitochondria than their midsegment counterparts. The spatial pattern of in situ pulmonary endothelial cell intracellular Ca2+ levels and reactive oxygen generation in response to TNF- stimulation positively correlate with the mitochondrial content at those sites (Parthasarathi et al. 2002). The major difference in the environment of midsegment and branch point cells is the magnitude and pattern of shear-stress. In the midsegment, flow is thought to be laminar and the shear-stress of consistent magnitude; while in the branch points, flow is thought to be turbulent and the shear-stress – highly variable. These observations suggest that shear-stress not only regulates the endothelial cell phenotype, but also affects the mitochondrial mass and their function in individual cells.

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Table 7.2. Mitochondrial morphological parameters Cell type and growth condition

PAEC in static culture PAEC flow-adapted in vitro PAEC in situ in rat lung PMVEC in static culture PMVEC flow-adapted in vitro PMVEC in situ in rat lung

Mitochondrial number/cell 257 145 36 167 95 24

± ± ± ± ± ±

43 26* 6 32 14* 7

Mitochondrial mass, μm3 404 199 35 557 279 19

± ± ± ± ± ±

34 18* 6 66 36* 5

Mitochondrial length, μm 8 7 5 17 15 4

± ± ± ± ± ±

3 2 2 6 4 2

Note:Mitochondrial mass was calculated from volume measurements, assuming a uniform cylindrical shape (diameter 0.5 m). The number of cells analyzed was 5–15. *p < 0.05 vs static culture of same cell type.

The most striking finding was the relative paucity of mitochondria compared to their content in endothelial cells in static culture. Endothelial cells in situ are naturally flow-adapted. Indeed, Table 7.2 reveals that flow-adapted cells in vitro and in vivo have less mitochondria, compared to their statically cultured counterparts. The decreased mitochondrial content may be related to decreased energy requirements of flow-adapted endothelial cells that are more quiescent than cells under static culture. Figures 7.3 and 7.4 show mitochondrial disposition in endothelial cells in static culture and in vivo (naturally flow-adapted) respectively. Mitochondria in naturally flow-adapted pulmonary precapillary endothelial cells in intact rat lung exhibit a paucity of mitochondria, all with an apparent perinuclear distribution. Average mitochondrial diameter of the flowadapted cells is approximately 0.5 μm. (Figure 7.3).

Figure 7.3. Intact organ fluorescence imaging of endothelial mitochondria in a subpleural pulmonary precapillary vessel in the rat lung. Isolated rat lung was perfused with 100 nM MitoTracker Green FM for 30 min and then the dye was washed out. Images of endothelial cell mitochondria in situ were acquired by transpleural imaging of the perfused lung with a 60x objective and Hamamatsu ORCA 100 ER digital camera using a Nikon 2000-TE microscope. Mitochondria appear as dots and thread and are located perinuclearally.

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Figure 7.4. Mitochondria in pulmonary microvascular endothelial cells were labeled with 100 nM Mitotracker Red 633, and then stacks of z-axis images were acquired at 0.5 μm intervals. A single projection from a 3-D reconstructed image was obtained after PSF-based deconvolution using MetaMorph (Molecular Devices, Inc) software.

In contrast to the endothelial cells in situ which contain a few mitochondria, rat pulmonary microvascular endothelial cells in static culture contain numerous mitochondria per cell with different shapes and sizes, which are scattered throughout the cytoplasm (Figure 7.5). However, when pulmonary microvascular cells were cultured in vitro under 20 dyn/cm2 shear-stress for 24 h, the cells assume the phenotype seen under in vivo flow. The mitochondria gradually acquire spherical shape, and after an initial increase, their velocity diminishes (Figure 7.6). These alterations indicate that the mitochondria may be an important site of mechanotransduction. This idea is also supported by the observation that disruption of microtubules, which interferes with mechanotransduction, prevents the maintenance of mitochondrial shape and structure (Figure 7.7). They remain straighter. This pattern of microtubule changes may alter the pattern of mitochondrial movement under flow, with the mitochondrial translocation being more restrictive when the microtubules assume the loop configuration (Figure 7.8). Kudo et al. (2000) have shown that endogenous ATP content increases 70% with acute imposition of shear-stress on statically cultured cells (Kudo et al. 2000). When endothelial cells were adapted first to 12 h of flow at 10 dynes/cm2

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Figure 7.5. Mitochondria in a pulmonary microvascular endothelial cell were labeled with 100 nM MitoTracker Green FM for 30 min and then the dye was washed out. Images were acquired with a 100x objective and Hamamatsu ORCA 100 ER digital camera using a Nikon 2000-TE microscope. Numerous mitochondria appear as elongated threads. The picture was acquired with a monochrome camera, therefore, no pseudocolor rendition was obtained.

Figure 7.6. Reduction in mitochondrial number and other morphological changes in a pulmonary microvascular endothelial cell under flow. Cells were plated on coverslips and subjected to 20 dyn/cm2 for up to 24 h in a parallel plate chamber. Pictures were taken after 2 h (left panel), 6 h (middle), and 24 h (right). Shape changes and gradual decrease in mitochondrial mass is observed with flow-adaptation. Labeling was with MitoTracker Green. The picture was acquired with a monochrome camera, therefore, no pseudocolor rendition was obtained.

Figure 7.7. Colchicine treatment led to disruption of mitochondrial structure and hazy labeling with MitoTracker Red 633, indicating loss of mitochondrial membrane integrity (bottom right). The picture was acquired with a monochrome camera, therefore, no pseudocolor rendition was obtained.

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Figure 7.8. Endothelial cells from rat pulmonary microvasculature (left panel) and pulmonary artery (right panel) were plated on coverslips and subjected to 20 dyn/cm2 for 3 h in a parallel plate chamber. Microtubules were stained with Oregon Green paclitaxel and mitochondria with MitoTracker Red 633.

in a dual-chamber parallel plate flow chamber, and then subjected to 6 h of non-hypoxic flow-cessation, ATP generation increased compared with statically cultured cells (Control, Figure 7.9). Hyperpolarization of mitochondria with flow-cessation (Figure 7.1) is compatible with the increased ATP generation. Twenty-four hour of flow-adaptation led to significant decrease in intracellular ATP levels compared to control (Figure 7.9).

Intracellular ATP levels, % of Control (Statically cultured EC's)

160 140

* - p 5μM, depresses cardiac function. Even a partial inhibition of CaUP may have a deleterious effect if the heart were stimulated with adrenaline and/or subjected to increased workload, assuming that an increase in [Ca2+ ]m under these conditions was essential for increasing ATP supply. However, to the best of our knowledge, this type of experiment has yet to be attempted. Similarly, use of inhibitors of the mNCX in intact cells and hearts is not without problems: of the 3 inhibitors that are effective in isolated mitochondria, namely diltiazem, clonazepam and CGP37157, we found only clonazepam to be effective in rat myocytes (Griffiths et al. 1997b). Diltiazem cannot be used because of its effects on L-type Ca2+ -channels, and we found that CGP37157 had variable effects on myocyte contractility and [Ca2+ ]m , possibly suggesting problems with cell permeability, or a non-mitochondrial effect, as in neuronal

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cells where CGP37157 was reported to inhibit voltage-gated Ca2+ channels (Baron and Thayer 1997). Clonazepam could be used as a relatively specific inhibitor of mNCX in adult rat myocytes, and at concentrations up to 100 μM had no significant effect on normal myocytes contraction or the cytosolic Ca2+ transient (Griffiths et al., 1997b). However, in adult guinea-pig myocytes, and in neonatal rat myocytes, clonazepam totally inhibited contraction (Bell & Griffiths, unpublished observations). A likely explanation for this is that clonazepam at these concentrations inhibits the sarcolemmal NCX, which is known to contribute to a much greater extent to Ca2+ removal from the cytosol during normal EC coupling in these cell types: in adult rat cells the SR accounts for over 95% of cytosolic Ca2+ removal (Bers 1991), so any effect of clonazepam on the NCX in these cells would go unnoticed.

9.2.5. Is There More than One Pathway for Ca2+ Influx? Whereas for many years there was thought to only be one Ca2+ -uptake pathway in mitochondria – the classical CaUP – more recently the existence of a rapid uptake mode (RAM) has been proposed (Buntinas et al. 2001; Sparagna et al. 1995). However, whether this represented a separate pathway or different state of the CaUP is unclear. Neither is it obvious how this pathway is reconciled with the original studies on isolated mitochondria, indicating that, although the CaUP can take up large amounts of Ca2+ , uptake is relatively slow. Another recent finding proposed the existence of a ryanodine receptor in the mitochondrial membrane (mRyR) (Beutner et al. 2001; Beutner et al. 2005); this appears to allow rapid uptake of Ca2+ at relatively low concentrations, and so could account for physiological beat-to-beat Ca2+ uptake. Since this pathway is inhibited as external [Ca2+ ] increases, it could have easily been missed in earlier studies on isolated mitochondria, the majority of which used relatively high external [Ca2+ ], and indicators that were not sensitive to sub-micromolar [Ca2+ ]. The authors suggest that as the ryanodine-sensitive pathway becomes inhibited, the classical CaUP takes over and accounts for mitochondrial Ca2+ uptake at higher external [Ca2+ ], and so would be the more pathologically relevant one (Beutner et al. 2005). These authors also found that ryanodine at concentrations of 2– 20 μM could inhibit large-amplitude swelling of mitochondria, a measure of MPTP opening (Beutner et al. 2005). < 2 μM ryanodine enhanced swelling, which was in agreement with its effects on the RyR-1 channel of skeletal muscle i.e. that at lower concentrations ryanodine activates the channel. It is still unclear as to whether the mRyR is the same as the CaUP – they are both inhibited by RuR and Mg2+ , suggesting they may be the same channel. But having two separate pathways is an attractive hypothesis, and would explain many of the controversies or discrepancies in results to date. For example, Ca2+ transients occur in adult guinea-pig and neonatal rat myocytes, but not in adult rat myocytes. Although highly speculative at present, a different expression of the Ca2+ transport pathways between species, and/or stage of development, could explain these results.

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9.3. Role of Mitochondrial Ca2+ in Ischaemia/ Reperfusion Injury 9.3.1. Alterations in [Ca2+ ]m During Ischaemia/ Reperfusion and Hypoxia/Reoxygenation Injury. Mitochondria have the capacity to take up huge amounts of Ca2+ (Gunter and Pfeiffer 1990; Nicholls and Crompton 1980) and thus could potentially remove toxic levels of Ca2+ from the cytosol. Unfortunately, such accumulation of Ca2+ can eventually damage mitochondria both by competing for ATP production and more importantly by inducing the mitochondrial permeability transition pore (MPTP). It has been known for many years that Ca2+ -induced mitochondrial dysfunction during ischaemia is associated with the transition from reversible to irreversible cell damage (Bush et al. 1980; Fleckenstein et al. 1983; Sordahl and Stewart 1980): Mitochondria accumulate deposits of calcium phosphate, become swollen, and eventually rupture. This sequence of events, whereby mitochondrial Ca2+ uptake leads to mitochondrial dysfunction (and hence prevents oxidative phosphorylation) was later found to be due to Ca2+ induced opening of the MPTP. The MPTP is discussed in detail in the chapter by Halestrap et al. in this volume. Similarly, there are several reviews detailing evidence for mitochondrial dysfunction in reperfusion injury (Di Lisa et al. 2003; Halestrap et al. 2004; Honda et al. 2005; Piper et al. 1994; Suleiman et al. 2001). In this chapter we will concentrate on the pathways of mitochondrial Ca2+ transport under pathological conditions, whether excessive mitochondrial Ca2+ uptake is beneficial or harmful under conditions of ischaemia/reperfusion, and whether the mitochondrial Ca2+ transport pathways could be targets for protective interventions. Mitochondrial Ca2+ uptake upon reperfusion of the heart has the potential to be protective, since it would effectively remove Ca2+ from the cytosol, and prevent cellular Ca2+ overload. Unfortunately, excessive Ca2+ uptake by mitochondria is damaging, firstly by inhibiting ATP synthesis (by lowering  m ), and secondly by inducing the MPTP. With regard to the first point,  m appears to remain constant during the contractile cycle in isolated myocytes; implying that any beat-to-beat changes in [Ca2+ ]m are too small to significantly affect  m under physiological conditions (Di Lisa et al. 1995; Griffiths 1999a), and so Ca2+ uptake would not be expected to affect ATP synthesis. In isolated mitochondria, respiration-driven Ca2+ uptake causes a dissipation of  m (Akerman 1978), which begins with external [Ca2+ ] as low as 1-2μM. However,  m recovers once Ca2+ uptake stops. With regard to induction of the MPTP by Ca2+ , heart mitochondria are more resistant than other tissues such as liver and kidney (Griffiths and Halestrap 1993), and can accumulate large amounts of Ca2+ , (20 nmol/mg protein, equivalent to approximately 10-20 μM) without triggering of the MPTP (Crompton et al. 1978). But if mitochondria are first depleted of adenine nucleotides, or subjected to oxidative stress, conditions that prevail upon reperfusion, then the MPTP is induced at a far lower [Ca2+ ]m (Griffiths and Halestrap 1995). The

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decrease in  m due to MPTP opening causes release of accumulated Ca2+ from mitochondria (Crompton and Heid 1978), a process inhibited by cyclosporine A (Crompton et al. 1988). [Ca2+ ]m increases upon reperfusion of whole hearts, and hypoxia/reoxygenation (or models of metabolic inhibition) of myocytes (Chacon et al. 1994; Delcamp et al. 1998), and Miyata et al. (1992) found that whether or not rat cardiomyocytes recovered from hypoxia depended on the level of [Ca2+ ]m achieved at the end of the hypoxic period: cells having [Ca2+ ]m greater than 250-300 nM invariably hypercontracted upon reperfusion (Miyata et al. 1992). This value of 300 nM [Ca2+ ]m is well within the normal physiological range of [Ca2+ ]m , and lower than that required for maximum activation of the dehydrogenase enzymes (0.5–1 μM). However other factors upon reperfusion can lower the threshold of [Ca2+ ]m needed for the MPTP to open; for example low adenine nucleotides and free radical generation. The values for increases in [Ca2+ ]m during hypoxia of isolated myocytes are in good agreement with those found in whole hearts: Allen et al (Allen et al. 1993) measured [Ca2+ ] in mitochondria isolated following hypoxia or reoxygenation of rat hearts perfused with fura-2. They found that 80 min of hypoxia caused no change in total tissue calcium, whereas free [Ca2+ ]m rose from pre-hypoxic values of 160 nM, to 360 nM and 570 nM after 50 and 80 min of hypoxia, respectively. Reperfusion after 50 min hypoxia resulted in no further change in [Ca2+ ]m , but after 80 min hypoxia, reperfusion led to a 10-fold increase in [Ca2+ ]m (Allen et al. 1993). This also highlights the fact that measurements of total tissue calcium cannot be extrapolated to measurements of free [Ca2+ ]. Mitochondria from different species also have different Ca2+ uptake capacities, which may influence their recovery from hypoxia/ischaemia; for example, the rate of Ca2+ uptake was greater in muskrat (a diving mammal) compared with guinea-pig mitochondria, and the former were also more resistant to Ca2+ induced MPTP opening (McKean 1991). The authors of this study suggested that this may aid the animal during recovery from hypoxia. Another factor to consider is the effect of gender: mitochondria from female rats had lower rates of mitochondria Ca2+ uptake than those from male rat heart, and the “female” mitochondria also had improved ability to repolarise following Ca2+ overload. It is well known that females are protected from some types of heart disease by oestrogen, and this may be one possible mechanism to explain the protective effects of oestrogen on ischaemia-reperfusion injury, although a mechanism for this has yet to be proposed (Arieli et al. 2004). But it should be remembered that limited (rather than excessive) Ca2+ uptake by mitochondria on reperfusion has the capacity to be protective: for example, by activation of pyruvate dehydrogenase (PDH). Several studies have shown that increased glucose oxidation on reperfusion improves recovery of the heart, and that drugs like ranolazine, which increase PDH activity, can improve postischaemic recovery (Clarke et al. 1993). So in designing protective strategies, it is a matter of getting the balance right between beneficial versus harmful effects of mitochondrial Ca2+ uptake. The situation is further complicated by the fact that many other factors on reperfusion affect this balance.

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9.3.2. Studies using Inhibitors of Mitochondrial Ca2+ Transport 9.3.2.1.

RUR and Ru360

RuR has been reported to protect hearts or myocytes from reperfusion/reoxygenation damage at concentrations ranging 0.1 – 6 μM (Benzi and Lerch 1992; Carry et al. 1989; Figueredo et al. 1991; Grover et al. 1990; Leperre et al. 1995; Miyamae et al. 1996; Park et al. 1990; Peng et al. 1980; Stone et al. 1989). Most of these studies have suggested that protection is due to effects of RuR on [Ca2+ ]m . However, few of these studies also reported effects on [Ca2+ ]c , and it seems more likely that the protective effects were due to an energy sparing effect as a result of reducing [Ca2+ ]c ; a conclusion reached by Benzi and Lerch (1992). We have found that much higher concentrations of RuR (about 20μM) are required to actually inhibit [Ca2+ ]m uptake in cardiomyocytes, and this concentration has a deleterious, not a protective, effect (Griffiths et al. 1998). So although low concentrations of RuR do appear to be cardioprotective, this cannot be attributed to direct effects on mitochondrial Ca2+ uptake. Ru360 has also been reported to protect whole hearts against ischaemia/ reperfusion injury: Pre-treatment of isolated rat hearts with 10 μM Ru360 provided protection against reperfusion injury, as determined from infarct size and enzyme release (Zhang et al. 2006). However another study reported that lower concentrations of Ru360 were required: functional recovery of rat hearts from ischaemia was optimal between 0.25–1 μM Ru360 (de Jesus Garcia-Rivas et al. 2005). Recovery declined at higher concentrations, and was not seen at 10 μM Ru360. This protective effect of Ru360 was maintained upon separation of mitochondria in terms of their ability to synthesise ATP and resist MPTP opening upon addition of external Ca2+ . [Ca2+ ]m (as measured following isolation of mitochondria at the end of the perfusion protocol) was also decreased in Ru360 treated hearts (de Jesus Garcia-Rivas et al. 2005). 9.3.2.2.

Inhibitors of the Na+ /Ca2+ Exchanger

Diltiazem can protect hearts from ischaemia/reperfusion damage, but this has mainly been attributed to its effects on sarcolemmal L-type Ca2+ -channels (Winniford et al. 1985); although a recent study suggested that diltiazem inhibits cytosolic Na+ overload through Na+ channels in the ischaemic heart, which would indirectly preserve mitochondrial integrity (Takeo et al. 2004). Clonazepam cannot be used in whole hearts as it appears to inhibit contractility by a non-myocyte effect (Griffiths, unpublished observation), and there have been no reports of the effects of CGP37157 in whole hearts. But clonazepam has been used in isolated myocyte models of hypoxia/reoxygenation injury: In suspensions of rat myocytes, 100 μM clonazepam reduced total cell [Ca2+ ], Ca2+ uptake (mitochondrial and cytosolic [Ca2+ ] were not measured separately) and LDH release, indicating preservation of membrane integrity (Sharikabad

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et al. 2001). Zhu et al. (2000) subjected cultured adult rat myocytes to 5mM extracellular Ca2+ , a condition that produced an increase in [Ca2+ ]m , measured using rhod-2, and in necrotic cell death (Zhu et al. 2000). Pre-treatment with clonazepam for 30 min reduced both the increase in [Ca2+ ]m , and the number of necrotic cells (to about 50% of control). Interestingly, cyclosporine A (CsA, an inhibitor of the MPTP) and clonazepam together reduced [Ca2+ ]m to similar levels as clonazepam alone, but the occurrence of necrotic cells was only 9% of control. This suggests that although clonazepam may have reduced [Ca2+ ]m in a sub-population of cells, the MPTP still opened in other cells, and presence of CsA then protected these cells from MPTP opening, and necrosis. Another study in isolated myocytes found that clonazepam (100μM) inhibited total cellular [Ca2+ ] increase in response to oxidative stress induced by H2 O2 (Sharikabad et al. 2004). As well as the Ca2+ transport inhibitors themselves, intracellular acidification, such as occurs during ischaemia, may also prevent excessive mitochondrial Ca2+ uptake: in isolated rat mitochondria, lowering external pH to 6.8 resulted in mitochondrial acidification – this was measured in permeabilised myocytes that had been previously loaded with the pH-sensitive fluorescent indicator carboxySNARF, under conditions where it localised into mitochondria (Gursahani and Schaefer 2004). The acid pH decreased the rate of Ca2+ uptake, probably as a consequence of reduced  m (Gursahani and Schaefer 2004) . Maintaining an acid pH upon reperfusion is known to delay intracellular Ca2+ accumulation and protect against reperfusion injury (Panagiotopoulos et al. 1990), and additionally inhibits opening of the MPTP (Halestrap 1991). 9.3.2.3.

Do the Transporters Reverse Direction During Hypoxia?

Although several studies had reported an increase in [Ca2+ ]m during hypoxia (see above), the mechanism of entry of the Ca2+ was either not discussed, or assumed to be the CaUP. But  m depolarises during hypoxia (Di Lisa et al. 1995), and this would be expected to inhibit Ca2+ uptake through the CaUP. In order to investigate the mechanisms and time-course of [Ca2+ ]m changes during cell injury, we used a model of single cell hypoxia/reoxygenation injury; this allowed continuous monitoring of [Ca2+ ]m , and correlation with cell functional recovery upon reoxygenation. We found that the increase in myocyte [Ca2+ ]m during hypoxia could not be prevented by RuR but instead could be prevented by clonazepam. In contrast, upon reoxygenation, RuR once again inhibited Ca2+ uptake, whilst clonazepam inhibited efflux (Griffiths et al. 1998). This allowed us to propose the following model: during hypoxia Ca2+ entry into mitochondria occurs via mNCX, and the CaUP is largely inactive. Upon reoxygenation, however, the transporters regain their normal directionality. This model of hypoxia/reoxygenation injury is represented in Figure 9.4. Clonazepam (but not RuR), also protected the cells from reoxygenation induced damage, as indexed by the myocytes’ ability to restore synchronised cell contractions (Griffiths et al. 1998). We have provided further evidence that entry of Ca2+ during hypoxia occurs via mNCX by showing, using a model of simulated hypoxia, that the

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E J. Griffiths, C.J. Bell, D. Balaska and G.A. Rutter NORMOXIA CaUP Ca2+

mNCX Ca2+ Ca2+ 3Na+

Ca2+

--- ++ Δψm negative (-180 mV) + +

HYPOXIA CaUP inactive

mNCX Ca2+ influx 2Na+ Ca2+

[Ca2+]m > 300 nM

[Ca2+]m < 300 nM

REOXYGENATION – SURVIVAL CaUP Ca2+ influx

mNCX Ca2+ efflux Ca2+ Ca2+

Δψm collapsed; CaUP inactive; mNCX reverses

3Na+

--

-++ + + Δψm restored; transporters regain normal directionality.

REOXYGENATION – CELL DEATH CaUP Ca2+ influx

mNCX Ca2+ efflux Ca2+ 3Na+

MPT P Δψm restored initially; but then MPTP is induced due to high [Ca2+]m.

Figure 9.4. Changes in cardiomyocyte mitochondrial Ca2+ transport pathways during hypoxia/reoxygenation injury – implications for cell survival. Studies using a single-cell model of hypoxia-reoxygenation injury found that the level of [Ca2+ ]m achieved at the end of hypoxia determined whether cells survived (recovered ability to respond synchronously to electrical stimulation), or hypercontracted (cell death). Cells only survived if [Ca2+ ]m remained below about 250-300 nM. The accumulation of [Ca2+ ]m during hypoxia also appeared to be via reversal of the mNCX, and not the CaUP (see text for further explanation). We proposed that upon reoxygenation, m initially repolarises. Then if [Ca2+ ]m remains below about 300 nM, ATP synthesis is restored, and the Ca2+ transport pathways of the cell regain their normal function. However if [Ca2+ ]m rises above about 300nM, this prevents ATP synthesis, and the MPTP is induced resulting in cell death.

process is Na+ -dependent (Griffiths 1999a). In support of this idea, a reversal of the mNCX under conditions of metabolic inhibition was also found in a study on cultured renal epithelial cells: CGP37157 inhibited the rise in [Ca2+ ]m (rhod-2 fluorescence) induced by metabolic inhibition (2-deoxyglucose plus cyanide) (Smets et al. 2004).

9. Mitochondrial Calcium

9.3.2.4.

213

Ischaemic Preconditioning

A discussion of the mechanism of ischaemic preconditioning (IPC) is beyond the scope of this chapter, but is dealt with elsewhere in this volume. However, one of the proposed protective mechanisms is that IPC leads to opening of a mKATP channel, and opening of this channel leads to membrane depolarisation (Murata et al. 2001). This, in turn, reduces Ca2+ uptake by mitochondria. A reduced [Ca2+ ]m has been reported in a model of IPC (Murata et al. 2001), but it is far from clear that this was the result of mKATP opening. Furthermore, the effect of, for example, diazoxide, on  m is controversial – some studies finding that it does produce a decrease (Murata et al. 2001), and others that it does not (Hanley et al. 2002; Katoh et al. 2002; Lawrence et al. 2001). The role of this mKATP channel has been almost entirely inferred indirectly based on studies using pharmacological modulation of the channel; by activators or inhibitors such as diazoxide and 5-hydroxdecanoate. An increasing number of studies recognise that these agents are not specific, and that their actions on the heart can be explained by actions other than on mKATP channels (Hanley et al. 2002; Katoh et al. 2002; Lawrence et al. 2001; Lim et al. 2002).

9.4.

Therapeutic Implications

As well as possible benefits of modulating [Ca2+ ]m during ischaemia/reperfusion injury, there is evidence that both RuR and Ru360 can reduce ventricular fibrillation (VF): In a pacing-induced model of VF in the isolated rat heart, perfusion with RuR (5μM), or Ru360 (10μM), resulted in conversion of VF to ventricular tachycardia (VT) (Kawahara et al., 2003). This is potentially beneficial since VT commonly occurs before VF, and sudden cardiac death (Myerburg et al. 1997). However, the benzodiazepines like clonazepam and diltiazem cannot be used in the whole heart as specific antagonists of mitochondrial Ca2+ efflux, because of their effects on coronary vessels (diltiazem, for example, is used to reduce high blood pressure). However, design of more specific inhibitors of the mNCX may be of benefit in states where ATP synthesis is impaired such as in cardiomyopathies, since maintaining [Ca2+ ]m at higher levels may in turn increase ATP production. There is some precedent for the idea that inhibiting the mNCX can enhance [ATP]: in pancreatic islets CGP37157 increased oxidative phosphorylation, and potentiated glucose-stimulated insulin release (Lee et al. 2003), prompting the authors to suggest it as a novel insulin secretagogue. An intriguing study looked at human mitochondrial complex I deficiency – this is associated with numerous clinical symptoms, ranging from lethal encephalopathy’s, of which Leigh disease is the most common, to neurodegenerative disorders, including Parkinson’s disease (Triepels et al. 2001). This study used skin fibroblasts from children with Complex I deficiency, and measured cytosolic Ca2+ transients in response to bradykinin (Visch et al. 2004). They found impaired increases in [Ca2+ ]c , [Ca2+ ]m , and ATP production in response to bradykinin in the complex I deficient fibroblasts. Treatment with 1μM CGP37157

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restored all these parameters. They suggested that (i) decreased ATP production in this disease is a result of reduced mitochondrial Ca2+ uptake, possibly due to a reduced  m in these patients, and (ii) CGP37157 can indirectly promote enhanced release of Ca2+ from the ER in these cells by enhancing mitochondrial Ca2+ uptake so that there is less inhibitory Ca2+ available at the mouth of the IP3 -operated ER Ca2+ release channels. Given the limitations on using the current mitochondrial Ca2+ transport inhibitors, it would be of great benefit if the transporters themselves could be cloned, for use in transgenic or knockout studies. Unfortunately there have only been a few sporadic attempts to purify the CaUP (Mironova et al. 1994; Saris et al. 1993; Zazueta et al. 1998) and mNCX (Li et al. 1992; Paucek and Jaburek 2004), and none has been more than partially successful. One of the main problems is that mitochondrial membrane proteins are notoriously difficult to purify, and the benefits of attempting their characterisation may require a lot of effort for little apparent gain. Nevertheless, a concerted effort to accomplish this may provide more benefit than can be foreseen at present.

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Trollinger DR, Cascio WE, Lemasters JJ (1997) Selective loading of Rhod 2 into mitochondria shows mitochondrial Ca2+ transients during the contractile cycle in adult rabbit cardiac myocytes. Biochem Biophys Res Commun 236(3):738–42. Tsien RY (1980) New calcium indicators and buffers with high selectivity against magnesium and protons: design, synthesis, and properties of prototype structures. Biochemistry 19(11):2396–404. Vasington FD, Gazzotti P, Tiozzo R, Carafoli E (1972) The effect of ruthenium red on Ca 2+ transport and respiration in rat liver mitochondria. Biochim Biophys Acta 256(1):43–54. Visch HJ, Rutter GA, Koopman WJ, Koenderink JB, Verkaart S, de Groot T, Varadi A, Mitchell KJ, van den Heuvel LP, Smeitink JA, Willems PH (2004) Inhibition of mitochondrial Na+-Ca2+ exchange restores agonist-induced ATP production and Ca2+ handling in human complex I deficiency. J Biol Chem 279(39):40328–36. White RL, Wittenberg BA (1993) NADH fluorescence of isolated ventricular myocytes: effects of pacing, myoglobin, and oxygen supply. Biophys J 65(1):196–204. White RL, Wittenberg BA (1995) Effects of calcium on mitochondrial NAD(P)H in paced rat ventricular myocytes. Biophys J 69(6):2790–9. Winniford MD, Willerson JT, Hillis LD (1985) Calcium antagonists for acute ischemic heart disease. Am J Cardiol 55(3):116B–124B. Ying WL, Emerson J, Clarke MJ, Sanadi DR (1991) Inhibition of mitochondrial calcium ion transport by an oxo-bridged dinuclear ruthenium ammine complex. Biochemistry 30(20):4949–52. Zazueta C, Zafra G, Vera G, Sanchez C, Chavez E (1998) Advances in the purification of the mitochondrial Ca2+ uniporter using the labeled inhibitor 103Ru360. J Bioenerg Biomembr 30(5):489–98. Zhang SZ, Gao Q, Cao CM, Bruce IC, Xia Q (2006) Involvement of the mitochondrial calcium uniporter in cardioprotection by ischemic preconditioning. Life Sci 78(7): 738–45. Zhou Z, Matlib MA, Bers DM (1998) Cytosolic and mitochondrial Ca2+ signals in patch clamped mammalian ventricular myocytes. J Physiol 507 ( Pt 2):379–403. Zhu LP, Yu XD, Ling S, Brown RA, Kuo TH (2000) Mitochondrial Ca(2+)homeostasis in the regulation of apoptotic and necrotic cell deaths. Cell Calcium 28(2):107–17.

10 Mitochondrial Ion Channels Brian O’Rourke

10.1.

Introduction

The maintenance of a large electrochemical driving force for protons across the mitochondrial inner membrane is essential for the production of ATP through oxidative phosphorylation. At face value, the opening of energy dissipating ion channels in the mitochondria would be unfavorable for energy transduction, but a wealth of evidence now indicates that selective (and some non-selective) ion channels may become active under various physiological or pathophysiological conditions. This review summarizes recent investigations into the functional roles for mitochondrial ion channels and efforts to identify molecular correlates related to specific ion fluxes across the inner membrane. While it is clear that mitochondrial ion channels play important roles in cellular life and death, our understanding of their structure is limited, and seminal discoveries are eagerly awaited. Interest in mitochondria has grown in recent years as the list of important cellular functions for this organelle has expanded from its fundamental role as a provider of energy to generator of reactive oxygen species (ROS), modulator of intracellular Ca2+ fluxes, initiator of cell death, mediator of cell protection, and participant in the processes of aging and disease. With regard to oxidative phosphorylation, the majority of the participating proteins have been identified at the molecular level, and in many cases, high resolution crystal structures are available. Importantly, a robust model of the energy transduction process, refined over the decades since the chemiosmotic principle was proposed, is available (for review, see Nicholls and Ferguson 2002). In contrast, many of the important proteins involved in mediating ion transport across the mitochondrial inner membrane have not been identified, although they have been well characterized at the functional level. Extensive evidence is available demonstrating that selective electrophoretic K+ (Brierley et al. 1971; Hansford and Lehninger 1972; Jung et al. 1977; Jung et al. 1984) and Ca2+ (reviewed by Gunter and Pfeiffer 1990) uptake by mitochondria occurs and that Na+ is exchanged for H+ (Douglas and Cockrell 1974) and Ca2+ (Jung et al. 1995). Similarly, K+ /H+ exchange balances the K+ uniport activity and modulates mitochondrial volume (Garlid 1996), yet no proteins have been 221

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definitively associated with any of these cation transport processes. Numerous transporters of anionic metabolites across the inner membrane, usually of the antiporter type, have also been identified at the molecular level (Jezek and Jezek 2003). However, apart from the inorganic phosphate carrier (Gerreira and Pedersen 1993) and the mitochondrial uncoupling proteins (UCP1, 2, and 3) (Rousset et al. 2004), little is known about the molecular structure of putative inner membrane anion channels that can mediate rapid anion flux. Since ion channels are capable of transporting millions of ions per second, and the electrochemical driving forces for ion movement across the inner membrane are enormous, the knowledge gap regarding mitochondrial ion channel structure is perhaps understandable – these proteins must necessarily be present in extremely low abundance, or have a very low open probability, in order to maintain the low permeability to ions required to exploit the protonmotive force for the generation of ATP via the mitochondrial ATP synthase. Nevertheless, although the opening of mitochondrial ion channels may be brief and highly controlled, the significance of the effects of their activation cannot be overstated. Mitochondrial ion channels are crucial to the mechanism of energy supply and demand matching and are the decisive factor in determining whether a cell lives and dies. Emerging evidence also suggests that the mitochondrial network can act as an intracellular signaling network. For example, the activation of redox sensitive transcription factors (e.g. HIF-1 or NF-B) and signalling pathways (e.g. protein kinase C) is likely to involve the generation of reactive oxygen species by the mitochondria (Guzy et al. 2005). Ion channels may contribute to this function in direct (serving as free radical transport pathways) or indirect (by altering the rate of respiration and the leak of electrons to ROS) ways. These novel and important functions of mitochondria underscore the need to identify and study the structure and organization of the proteins in the mitochondrial outer and inner membranes. While much progress has been made in assaying the function of mitochondrial ion channels and exchangers, the challenge ahead will be to assign structures to the various ion transport pathways and to study how increasing or decreasing their expression affects the integrated cell function.

10.2.

Fast Ion Movements Across the Inner Membrane

A basic tenet of the chemiosmotic hypothesis is that, apart from the movement of protons through the mitochondrial ATP synthase and through non-specific low conductance “proton leak” pathways, the mitochondrial inner membrane must be quite impermeable to ions in order to maintain the efficiency of oxidative phosphorylation. However, the large electrical driving force for ion movement (∼180mV) would strongly favor ion flow through any open ion channels, even in the absence of a concentration gradient across the inner membrane. A classic and unquestionable example is Ca2+ influx through the Ca2+ uniporter. Under conditions that prevent (e.g. in the presence of nucleotides) the activation of the mitochondrial permeability transition pore (PTP), mitochondria can take up enormous amounts of Ca2+ (up to almost 1M total Ca2+ ) although the matrix free

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Ca2+ concentration remains in the M range due to the reversible precipitation of Ca2+ with Pi. This Ca2+ sink property of the mitochondria is circumvented if PTP opening is triggered by the Ca2+ overload, allowing both the release of the accumulated Ca2+ and other matrix constituents with mass